Abstract
Uridine is the ubiquitous nucleoside form of the RNA base uracil. It occupies a prominent ‘hub’ position in energy metabolism; for example, it is metabolically linked to de novo pyrimidine biosynthesis and glycolysis and biologically linked to diverse processes, such as RNA synthesis/degradation and glycosylation. It is a vital interorgan ‘currency’ nutrient readily imported by mammalian cells, and its supplementation can exert both cytoprotective and toxic effects, for which the underlying mechanisms are poorly understood. Importantly, it is a route by which the decay of RNA can be repurposed as an alternative fuel source under nutrient-limiting conditions to aid in tumor initiation, development and metastasis. Here we explain how the upstream inputs and downstream metabolic fates of uridine influence cancer traits and illustrate both established and hypothetical strategies targeting uridine metabolism for cancer therapy.
Similar content being viewed by others
Overview of uridine
Uridine is a nucleotide that is common to all life forms and occupies a unique and complex position in metabolism from both an intracellular and intercellular perspective. In the intracellular metabolic network, it is adjacent to uridine monophosphate (UMP) and thus it can either be formed via the de novo pyrimidine biosynthetic pathway or, conversely, provide UMP in cells lacking it (Part 2). Thus, uridine metabolism is intertwined with key biological processes such as DNA/RNA synthesis, particularly RNA synthesis, as it uses the uridine base, as well as glycosylation, which utilizes uridine diphosphate (UDP) as the sugar carrier molecule (Fig. 1). Furthermore, its catabolism can support both glycolysis and the tricarboxylic acid (TCA) cycle (Part 3).
Uridine can be directly taken up from plasma, produced via the pyrimidine biosynthesis pathway, or obtained from RNA decay. It can then contribute to RNA/DNA synthesis, the TCA cycle, glycosylation and glucuronidation. Figure created using Biorender (https://biorender.com/).
From an intercellular/organismal perspective, uridine is also an important interorgan nutrient. Uridine, the natural nucleoside precursor of UMP, is reported to exist in human plasma at 3–8 μM, which is substantially greater than other nucleosides1,2,3, and is predominantly produced by the liver or adipose tissue for utilization across other tissues4. In addition, in human serum, UMP can be rapidly dephosphorylated to uridine in the presence of 5′-nucleotidase5,6. Thus, cells can readily obtain uridine from the environment, and the administration of uridine is cytoprotective in a wide range of contexts, including chemotherapy and ischemia7,8,9. On the other hand, the disruption of uridine homeostasis by the knockout of uridine phosphorylase can cause spontaneous tumors in mice and uridine treatment of cells can cause DNA damage and toxicity, suggesting that uridine homeostasis is essential10,11.
All of these aspects have implications for cancer cells and here we review the upstream metabolic factors that lead to uridine formation and the downstream impact of uridine metabolism from the perspective of cancer cells. We discuss how cancer cells obtain uridine, what they use uridine for and how any of these steps may be targeted for cancer therapy.
Part 1: uridine is a metabolic ‘currency’
While most metabolites, such as nicotinamide adenine nucleotide hydrogen and glycolytic intermediates, are produced and metabolized intracellularly, some such as glucose and ketone bodies (acetoacetate, 3-β-hydroxybutyrate and acetone), represent organism-level metabolic ‘currencies’—metabolites that are regulated at the whole-organism level, widely circulated and provided through the plasma to a variety of cell types12,13,14,15. For example, multiple organs in the body utilize glucose and regulate its circulatory levels14,16. In the same manner, uridine in the plasma is freely available for uptake by cells and uridine abundance is tightly regulated through various mechanisms, particularly via synthesis and degradation in the liver17,18 (Fig. 1). Under normal-fed states, the liver is the primary source of uridine biosynthesis as well as catabolism in the body4. However, under fasted states, pyrimidine biosynthesis from adipose tissue results in compensatory overproduction of uridine4. In addition to fasting, various physiological factors, such as alcohol consumption, diet and exercise, strongly regulate plasma uridine levels17,19,20. For example, uridine levels can be elevated with the consumption of certain uridine-rich foods, such as beer, or via ethanol itself19,21.
Uridine, in turn, is readily taken up by cells, as demonstrated in an organism context and in cell/tissue culture systems. Uridine uptake is mediated by both equilibrative nucleoside transporters (ENTs) and concentrative nucleoside transporters (CNTs), which can deliver a range of nucleosides (ENTs and CNTs)22 (Fig. 2), and the injection of supraphysiologic uridine can result in rapid uptake in most tissues, especially renal tissues23. Uridine uptake in various cell lines is evident24,25, and radiolabeled uridine uptake and incorporation into RNA has even been used as a measure of cell viability26.
Uridine can be imported into cancer cells via CNTs and ENTs to competitively inhibit the incorporation of 5-FU into RNA, minimizing gastrointestinal toxicity while retaining its cytotoxicity to cancer cells. Uridine also has various cytoprotective properties and suppresses oxidative stress and inflammation. ROS, reactive oxygen species. Figure created using Biorender (https://biorender.com/).
Uridine uptake in cancer
Intriguingly, in stark contrast to glucose, whose uptake and downstream utilization have been central points of focus in cancer metabolism research27,28, relatively little is known about the characteristics of uridine uptake and utilization by cancer cells. In particular, there is no consensus on whether certain cancers have elevated uridine uptake and for what purpose; however, much on uridine uptake can be extrapolated from studies on nucleoside analog uptake in tumors, as nucleoside transporters are also expected to mediate uridine uptake. In fact, labeled uridine uptake is often used as a proxy for the nucleoside analog uptake capacity of tumors29,30. Similarly, the overexpression of specific nucleoside transporters has been linked with tumor sensitivity to nucleoside analogs, and the loss of transporter expression has been linked with resistance31,32,33,34,35. However, these studies did not directly compare the uridine uptake of tumor tissues with that of normal tissues; thus, it remains an open question whether certain subtypes of tumors may have increased uridine uptake and be particularly dependent on uridine as a metabolic currency.
Consequences of uridine uptake
Uridine exerts various activities across organs, supporting its role as a currency metabolite. For example, its administration regulates reproductive organ function, vasoregulation and neuronal activity36,37,38, demonstrating that in normal physiology, uridine provided from the plasma is indeed of consequence to various cells. In one of the best-documented and cancer-related applications, uridine has been used in forms such as uridine triacetate as a cotreatment to reduce patient toxicity during nucleoside chemotherapy, such as with 5-fluorouracil (5-FU), while maintaining antitumor toxicity7,39. It is postulated that the protective effect occurs by competitively limiting the uptake of 5-FU into RNA, which is a major source of gastrointestinal toxicity40 (Fig. 2). Uridine has even been identified as a potent inducer of tissue regeneration41. Uridine supplementation has been successfully implemented for rare disorders such as orotic aciduria and carbamoyl phosphate synthetase II, aspartate transcarbamylase and dihydroorotase (CAD) deficiency, which are disorders in which enzymes responsible for pyrimidine biosynthesis, such as uridine monophosphate synthetase (UMPS) or CAD, are mutated; thus, uridine provides alternative precursors for pyrimidine synthesis42,43.
Intriguingly, uridine has various effects on cells in tissue culture. For example, it is cytoprotective in various conditions of stress, such as rescuing viability against oxidative phosphorylation or mitochondrial deficiency44,45, glucose deprivation8, inflammation and oxidative stress9. In oxidative phosphorylation or mitochondrial deficiency, uridine supplementation provides alternative precursors for pyrimidine biosynthesis, which are needed as the pyrimidine biosynthetic enzyme dihydroorotate dehydrogenase (DHODH) is dependent on the electron transport chain46. However, there may be additional undiscovered mechanisms of uridine-based cytoprotection, such as during oxidative stress. On the other hand, high levels of uridine can negatively affect cell viability, such as by inducing ferroptosis in hepatocellular carcinoma cells11 and causing DNA damage and p53 activation in a process referred to as ‘uracil damage’10. The same study revealed that the loss of uracil processing via uridine phosphorylase, resulting in uridine hyperaccumulation, can be carcinogenic. However, the aforementioned successful use of uridine supplementation in protecting against chemotherapy-induced toxicity or in-born metabolic disorders, without any noted side effects42,43, suggests that, in moderation, uridine can be safe and beneficial.
Thus, it is clear that uridine is a currency metabolite with a wide range of consequences for the cells that access it. To understand the underlying mechanisms of the effects of uridine in cancer cells and explore future options for therapeutic exploitation, it is important to review what is known about the downstream metabolic fates of uridine metabolism, as outlined below. Additionally, we consider other sources of uridine aside from the extracellular environment, particularly the de novo pyrimidine biosynthesis pathway, where the pathway end-product UMP is interconverted with uridine. Additionally, we consider nucleic acids themselves, particularly RNA, as sources of uridine upon their decay. In the sections below, we explore the various connections of uridine with other metabolic pathways, the potential importance of uridine from the perspective of cancer cells and how these connections may be potential points of vulnerability.
Part 2: upstream contributors to uridine metabolism
UMP as a source of uridine and vice versa
The metabolic fates of uridine and UMP are intertwined and the two molecules are only a bidirectional metabolic step removed from each other. Uridine, obtained from plasma, can be converted to UMP via uridine cytidine kinases (UCKs). On the other hand, UMP can be biosynthesized via a pathway that utilizes glutamine and bicarbonate to produce carbamoyl phosphate, followed by the incorporation of aspartate to generate carbamoyl aspartate as the initial step47,48. It can also be obtained from the breakdown of RNA and then converted to uridine via 5′ nucleotidases24. As described in the following sections, the relationship is bidirectional and fuels various requirements of cancer cells, including glycolysis occurring downstream of uridine or RNA synthesis and glycosylation occurring downstream of UMP.
Uridine to UMP via UCKs
Uridine can be phosphorylated to UMP via UCK, UCK1 and UCK2 in humans49,50 (Fig. 3). UCK1 is present in both cancer and normal cells, whereas UCK2 is preferentially expressed in various cancer cells, including neuroblastoma cells51,52,53. Given that uridine obtained extracellularly can contribute to UMP processes, radiolabeled uridine is readily incorporated into RNA26 and uridine administration induces glycosylation in cells and in vivo36,54.
Figure created using Biorender (https://biorender.com/).
UCK enzymes have been explored in cancer therapy, mainly for their role in the metabolic activation of ribonucleoside analog chemotherapeutics. UCK phosphorylates the ribonucleoside analog TAS 106 (3′-ethynylcytidine) to generate 3′-ethynylcytidine-5′-monophosphate51 that undergoes two phosphorylation steps to be converted into 3′-ethynylcytidine-5′-triphosphate, which inhibits RNA polymerases and exerts an antitumor effect51. TAS 106 showed excellent antitumoral activity in HT-1080 (half-maximum inhibitory concentration (IC50) of 0.01 μM) and NUGC-3 (IC50 of 0.35 μM) cells51. Notably, the level of UCK2, rather than that of UCK1, is closely correlated with the sensitivity of various cancer cells, including pancreatic cancer cells, to TAS 106 (ref. 55). The cytidine analog fluorocyclopentenylcytosine, RX-3117, is activated by UCK2, and its di- and triphosphorylated metabolites are incorporated into RNA and DNA, inhibiting gene synthesis56,57,58. RX-3117 exhibited potent antitumor activity in MDA-MB-231 (IC50 of 0.18 μM), NCIH226 (IC50 of 0.25 μM) and HCT116 (IC50 of 0.19 μM) cells59,60. Notably, UCK1 knockdown by short interfering RNA did not affect the sensitivity of A549 and SW1573 cells to RX-3117, whereas UCK2 knockdown reduced the sensitivity of A549 and SW1573 cells to RX-3117 (ref. 61). While such roles of UCK enzymes are well known, less is known about the implications of blocking the uridine-to-UMP flux that would occur upon UCK disruption, which is a potential avenue for therapy.
In addition to its catalytic role as a UCK, UCK2 has recently been shown to play important catalysis-independent roles, particularly in activating oncogenic signaling pathways such as the EGFR‒Akt axis through direct binding62,63. Thus, in addition to catalytic inhibitors, strategies that target UCK2 via degradation constitute an intriguing therapeutic avenue.
Glutamine to UMP via the pyrimidine biosynthesis pathway
The initial three reactions of the UMP de novo synthesis pathway, also known as the pyrimidine biosynthesis pathway, are catalyzed by CAD, generating dihydroorotate from bicarbonate, glutamine and aspartate64,65 (Fig. 4). Dihydroorotate is processed to orotate by DHODH on the outer face of the inner mitochondrial membrane, and orotate is metabolized in two steps, which are both catalyzed by the bifunctional enzyme UMPS66,67. The N-terminal domain of UMPS first catalyzes the conversion of orotate to orotidine monophosphate (OMP), utilizing phosphoribosyl diphosphate (PRPP) as a cosubstrate. The C-terminal domain of UMPS then catalyzes the conversion of OMP into UMP. Interestingly, the activity of this pathway is low in nonproliferating (that is, most normal) cells, whose demand for pyrimidines is largely met by salvage pathways68. On the other hand, this pathway is generally highly active in proliferating cells, which have high pyrimidine demand69. This finding raises the possibility that UMP produced in cancer cells can feed into uridine production. Conversely, owing to the high pyrimidine demand for RNA/DNA synthesis, uridine uptake by cancer cells may feed into the formation of UMP and subsequent formation of pyrimidine, which is used to fuel cancer growth. Future research, such as using labeled glutamine as a precursor for UMP formation, is needed to elucidate the exact relationship between uridine uptake and the pyrimidine biosynthesis pathway in cancer cells.
The UMP produced by this pathway can be converted to uridine via 5′-nucleotidases. Figure created using Biorender (https://biorender.com/).
Among the enzymes involved in de novo synthesis pathways, the expression and activity of DHODH are reported to be directly associated with cancer progression70. Arava (leflunomide), an inhibitor of DHODH, received United States Food and Drug Administration approval in 199871, and its active metabolite aubagio (teriflunomide) received Food and Drug Administration approval in 201271. DHODH, in addition to its role in pyrimidine biosynthesis, has received attention for its role in suppressing ferroptosis72. However, the observed effects of the DHODH inhibitor brequinar on sensitization to ferroptosis may be due to the inhibition of another enzyme, FSP1 (ref. 73); thus, further efforts are warranted to clarify the roles of DHODH in cancer cells.
Cytidine salvage to uridine
Uridine can be formed through the transformation of cytidine salvaged from the bloodstream74. This process is dependent on cytidine deaminase (CDA), which converts cytidine to uridine. CDA has been reported to be resistant to cytidine analogs, including cytosine arabinoside and 2′,2′-difluorodeoxycytidine (gemcitabine), since CDA metabolizes cytidine analogs into inactive metabolites75,76. CDA inhibitors have been developed, but primarily explored for their ability to negate the resistance-promoting property of CDA75,76. Whether CDA is relevant to cancer cells as a significant source of uracil, relative to other sources described, remains an interesting research direction and potential application for CDA inhibitors.
Part 3: RNA decay as an intracellular source of uridine
In eukaryotes, most messenger RNAs are protected from degradation by a 5′-7-monomethyl guanosine (m7G) cap and a 3′-poly(A) tail77,78,79. RNAs are also reported to undergo distinct noncanonical 5′-end modifications, including nicotinamide adenine dinucleotide and dephospho-CoA80,81. RNA decay is initiated by the removal of the 3′-poly(A) tail, and deadenylated RNA can be degraded through two pathways: 3′-to-5′ RNA decay and 5′-to-3′ RNA decay82,83. RNA is a highly abundant molecule in cells, constituting 4–20% of dry mass84. mRNAs turn over faster than other types of RNA85, and their decay and subsequent generation of UMP may be a substantial contributor to uridine metabolism86. In addition, given that ribosomal RNA is the most abundant type of RNA, ribophagy resulting in rRNA degradation in vacuoles is a major source of nucleotides during nutritional stress87. In support of the notion of RNA as a major source, uridine production derived from RNA is efficient enough to allow the growth of cancer cells in the complete absence of glucose24. In the following section, we discuss what is known about the role of RNA decay processes in cancer (Fig. 5). These previous studies have elucidated how various cellular processes are modulated by these RNA decay pathways in cancer and implicated various parts of the decay machinery as potential cancer targets. We propose that these targets may have additional relevance to cancer metabolism in light of RNA being a rich source of uridine, a concept that deserves further exploration.
The 3′-to-5′ decay is initiated by removal of the poly(A) tail by the CNOT complex, then decay can occur in RNA exosomes and the remaining 5′ cap is processed by DcpS. The 5′-to-3′ decay involves an initial decapping by the DCP1A/2 complex and then processing by the exonuclease XRN1. Viral-induced RNA decay involves the degradation of double-strand viral RNA by OAS enzymes, which results in 2′-5′ oligoadenylate formation, which is in turn further degraded by RNase L. In all of these cases, nucleoside monophosphates are formed, through which UMP can be directly converted to uridine. CMP can also be metabolized to UMP, providing a further source of uridine. AMP, adenosine monophosphate; CMP, cytidine monophosphate; GMP, guanosine monophosphate. Figure created using Biorender (https://biorender.com/).
The 3′-to-5′ RNA decay
Shortening and removal of the 3′-poly(A) tail of mRNA are the first steps of RNA decay, and this deadenylation is catalyzed by the deadenylase complexes PAN2–PAN3 and CCR4–NOT (CNOT)88. PAN2–PAN3 shortens the initial 3′ poly(A) tail, and CNOT completes the remainder of the deadenylation89. The CNOT complex consists of nine core subunits CNOT1, CNOT2, CNOT3, CNOT6, CNOT6L, CNOT7, CNOT8, CNOT9/RQCD1 and CNOT10 (ref. 90). Various CNOT subunits have been implicated in different cancer contexts. CNOT2 is implicated in breast cancer and colorectal cancer cell viability91,92, and its inhibition sensitizes non-small-cell lung cancer cells to TNF-related apoptosis93. Similarly, CNOT3 has been implicated as a driver of leukemogenesis by promoting translation efficiency94 and as a mediator of chemoresistance in lung cancer95,96. CNOT7 is involved in breast cancer metastasis97. The modulation of various signaling pathways, including the mTOR, STAT3, VEGF and c-Myc pathways, is associated with these effects.
Following deadenylation, mRNA can be degraded in the 3′-to-5′ direction by cytoplasmic multisubunit RNA exosome complexes98. The 5′ cap on the remaining oligomer is hydrolyzed by the decapping scavenger enzyme (DcpS), which converts the 5′ cap into 7-methylguanosine monophosphate (m7GMP) and nucleoside diphosphate98,99,100,101,102. DcpS has been shown to play an important role in mRNA homeostasis and to protect cells from the potential toxic accumulation of capped mRNA fragments102. DcpS enzymes, which form homodimers with two active sites situated in the grooves between the N- and C-terminal domains, can bind two cap ligands103,104. DcpS has recently been identified as an essential gene for acute myeloid leukemia (AML) cell survival through genome-wide CRISPR–Cas9 knockout screening. Depletion of DcpS by RG3039 or short hairpin RNA imparts potent antileukemic activity in human AML cells105,106. In addition, JCS-1, an E3 ligase VHL-recruiting PROTAC, has been developed to promote the degradation of DcpS, which inhibits the proliferation and viability of MOLM-14 cells107. These studies indicate that targeting DcpS could be an effective strategy for treating AML- and DcpS-dependent diseases.
The 5′-to-3′ RNA decay
Unlike 3′-to-5′ decay, Dcp2 removes the m7G mRNA cap in 5′-to-3′ decay108,109,110,111, and decapped RNA is rapidly degraded by the 5′-to-3′ exonuclease XRN1 (refs. 82,112,113). XRN1 has recently been implicated as a cancer target whose disruption potentiates the effect of cancer immunotherapy via a mitochondrial antiviral-signaling protein-dependent mechanism114,115. The metabolic implications of targeting XRN1 should be explored in relation to immunotherapy, considering that targeting nucleotide metabolism could enhance cancer immunotherapy116.
Endoribonuclease-mediated RNA decay
The 2′,5′-oligoadenylate (2–5A) synthetase (OAS)-–ribonuclease L (RNase L) pathway is a key component of antiviral innate immunity, which induces the degradation of viral and cellular RNAs, thereby blocking viral infection117. The antiviral 2–5A pathway is initiated by the binding of viral double-stranded RNA (dsRNAs) to 2′-5′-oligoadenylate synthetases, and activated OASs produce 2′-5′-linked oligomers called 2–5A [px5′A(2′p5′A)n; x = 1 − 3; n ≥ 2], with ATP used as a substrate118,119,120. The human OAS gene family consists of three functional genes, OAS1, OAS2 and OAS3, as well as the OAS-like gene, which encodes a protein that does not synthesize 2–5A121. The transcription of OAS is induced by interferon (IFN) signaling, and elevated levels of OAS contribute to the IFN-induced antiviral state117,122. Notably, compared with normal prostate epithelial cells, prostate cancer cells (PC3, LNCaP and DU145 cells) overexpress mRNAs encoding proteins that bind and activate OAS, such as Raf kinase inhibitor protein (RKIP), poly(rC)-binding protein 2 (PCBP2) and human endogenous retrovirus (hERV) envelope RNAs123,124. This finding indicates that OAS could be activated independently of virus infection in cancer.
RNase L, known as an IFN-induced endoribonuclease, is activated by 2–5A produced by OAS, resulting in the cleavage of all RNAs in the cell123,125. RNase L suppresses the mobile genetic element LINE-1 and stimulates apoptosis, inflammation and autophagy123,126,127,128,129. RNase L knockdown by shRNA increases the migration of both human prostate cancer cells and mouse embryonic fibroblasts123. In addition, the overexpression of RNase L inhibits androgen receptor signaling and migration in prostate cancer cells130. Considering these findings, the mutation status of the RNase L gene might be a key factor in promoting cell migration and metastasis in prostate cancer. Notably, IFN-γ enhances the expression of RNase L and restores its function in the cytoplasm and nucleus, which leads to apoptosis in lung cancer cells131. These findings indicate that RNase L could act as an adjuvant to enhance the efficacy of cancer immunotherapy via IFNs such as IFN-γ.
On the basis of these various potential sources of uridine for cells, we propose the following model for how cancer cells within solid tumors obtain uridine (Fig. 6). In tumor cells that are proximal to working vasculature, uridine should be readily obtained from the blood supply (Fig. 6, left). They may additionally obtain uridine from the pyrimidine biosynthesis pathway, which is upregulated in proliferating cells to meet the nucleic acid demands of a growing cell132. However, in a poorly vascularized tumor or a cancer cell that is distal to the blood supply, it is likely that not only glucose but also precursors (for example, aspartate and glutamine) for pyrimidine biosynthesis are highly limited. Therefore, recycling from the abundant RNA molecules in the cell is probably a critical source of uridine and downstream products in these cells. This model is suggested based on recent studies showing that exogenously provided uridine or RNAs can provide an alternative fuel source under glucose limitation24,25. However, further studies are needed to directly examine the importance of RNA decay in providing uridine as an alternative fuel source in the starved tumor microenvironment.
The upstream sources of uridine may depend on the availability of uridine itself or various precursors and, in states of depletion, RNA decay may provide a key source of uridine. Figure created using Biorender (https://biorender.com/).
Part 4: downstream fates of uridine to fuel cancer cell processes
Uridine contribution to glycolysis and the PPP
Recently, uridine was shown to be a key contributor to central carbon metabolism, particularly when glucose is limited for tumor cells. Uridine can be converted into ribose 1-phosphate and uracil by uridine phosphorylase (UPP)25, and it can be phosphorylated to produce UMP via UCK49,50,133. The removal of the phosphoryl group from UMP by 5′-nucleotidase results in uridine, and the conversion of UMP to OMP is mediated by orotidine 5′-phosphate decarboxylase in de novo synthesis134. Recently, it has been reported that ribose derived from uridine by UPP1 can overcome the growth defects induced by glucose deficiency in pancreatic cancer and that high UPP1 expression predicts poor outcomes in patients with pancreatic cancer24,25. Thus, uridine metabolism can be considered a source of fuel for cancer proliferation, emphasizing its potential as an anticancer target (Fig. 7).
The pathways and drugs that can target specific steps are outlined. Figure created using Biorender (https://biorender.com/).
Uridine to uracil and ribose 1-phosphate by UPP
Glucose, a primary carbon and energy source for cell growth, proliferation and activity, is metabolized through multiple pathways, including glycolysis and the pentose phosphate pathway (PPP)135. The increased uptake and dependency of cancer cells on glucose and their tendency to secrete carbon as lactate even under aerobic conditions were key early observations in the cancer metabolism field136. These properties are exploited in tumor imaging via 18F-fludeoxyglucose–positron emission tomography137. Blocking the uptake or metabolism of glucose is considered a strategy for inhibiting tumor growth138,139,140. However, an insufficient glucose supply is a situation commonly encountered by solid tumors, and cancer cells can utilize nutrients such as amino acids, lactate, acetate and other macromolecules as alternative fuels141.
Recently, uridine has been shown to serve as an alternative fuel for cancer growth in the absence of glucose24,25. In this process, uridine is converted to uracil and ribose 1-phosphate by UPP. In vertebrates, there are two forms of the phosphorylase, UPP1 and UPP2 (ref. 142). UPP expression and activity are induced by redox or chemotherapeutics; UPP2 activity is affected by the redox state due to a disulfide bridge, and the KRAS–MAPK pathway and NRF2 can modulate UPP1 expression25,142,143. The steady-state level of UPP1 mRNA is positively correlated with the poor prognosis of patients with multiple types of malignant tumors, including breast cancer, lung adenocarcinoma and oral squamous cell carcinoma144,145,146. Owing to the role of UPP1 in uridine usage under glucose-limiting conditions, how UPP1 is regulated and its implications for tumor outcome remain important topics for continued investigations.
Targeting UPP enzymes has previously been shown to impair various functions of cancer cells, which may in part include the interaction of UPPs with other proteins. UPP1 knockdown by siRNA significantly suppresses the migration, invasion and proliferation of TPC and BCPAP thyroid carcinoma cells147. Similarly, the pharmacological inhibition of UPP1 using CPBMF65, a synthetic human uridine phosphorylase-1 inhibitor, reduces the proliferation of HepG2 cells, leading to cell cycle arrest and senescence148. In lung adenocarcinoma, UPP1 knockdown via shRNA reduced the protein levels of ENO1 and LDHA in H292 and H1975 cells, whereas overexpression of UPP1 restored the suppression of lactate production, glucose uptake and ENO1/LDHA protein levels induced by 2-deoxy-d-glucose in H1299 cells145. A specific inhibitor of UPP, 5-(phenylthio)acyclouridine (PTAU), attenuated the reduction in glucose deprivation-mediated cell death caused by uridine in lipopolysaccharide- and IFN-γ-treated astrocytes149. In addition, UPP1 binds the C-terminus of AKT and increases the activity of AKT by promoting the interaction of AKT with PDK1 and PDK2 and recruiting phosphatidylinositol 3,4,5-triphosphate to AKT150. This is related to tumorigenesis and resistance to the drug gemcitabine in bladder cancer150. UPP2 has the potential to sense and initiate cellular responses to oxidative stress142. In addition, hepatic nuclear receptor agonists have been reported to regulate mouse UPP2, implying an additional function of UPP2 in hepatic lipogenesis or cholesterol transport142,151,152. In summary, multiple metabolic and signaling mechanisms justify UPP1 as an attractive target for anticancer treatment strategies.
Ribose 1-phosphate to R5P by PGM2
Phosphoglucomutase (PGM) catalyzes the reversible conversion between glucose 6-phosphate and glucose 1-phosphate using Mg2+ (refs. 153,154,155). The PGM protein family members PGM1–5 share homologous coding sequences, but their substrates and functions vary154. In the presence of uridine and the absence of glucose, phosphoglucomutase 2 (PGM2) connects the UPP1/UPP2 reaction to the PPP by catalyzing the conversion of ribose 1-phosphate to ribose 5-phosphate (R5P)24. Skinner et al. reported that PGM2 is essential for survival and proliferation in glucose-deprived, uridine-rich media, but is dispensable in glucose-containing media, indicating its essential role in the utilization of uridine as an energy source in the absence of glucose24. Additionally, PGM2 mRNA accumulates in cells of various cancers (lung, central nervous system, pancreas and small/large intestine), and this is associated with an unfavorable prognosis in patients with lung adenocarcinoma156. Considering these findings, targeting PGM2 could effectively inhibit uridine metabolism and be utilized as a target in treating cancer.
R5P to glyceraldehyde 3-P by TKT
Transketolase (TKT), a key enzyme of the nonoxidative PPP, catalyzes the transfer of two carbons from xylulose 5-phosphate to the co-factor of TKT, thiamine pyrophosphate, which then delivers them to R5P or erythrose 4-phosphate. R5P and erythrose 4-phosphate are converted into sedoheptulose 7-phosphate (S7P) and fructose 6-phosphate, respectively, and xylulose 5-phosphate is converted into glyceraldehyde 3-phosphate157,158. Glyceraldehyde 3-phosphate is subsequently converted into pyruvate through the second phase of glycolysis, which is then converted into acetyl-CoA, allowing it to enter the TCA cycle for ATP production159,160. TKT is transcriptionally regulated by various biomolecules, including HIF-1α, miR-497 and steroid receptor coactivator-3 (refs. 157,161,162,163,164,165). The transcription of TKT in imatinib-resistant chronic myelogenous leukemia cells is regulated by HIF-1α, sustaining the survival and proliferation of these resistant cells166. Transfection of miR-497 mimics targeting TKT mRNA increases the sensitivity of HeLa and SiHa cells to cisplatin, increasing their viability164. PFKFB4 promotes TKT transcription by activating oncogenic steroid receptor coactivator-3, which enhances breast tumor growth and metastasis in immunocompromised mice165.
Furthermore, TKT is overexpressed in various cancers, including colorectal, pancreatic, breast and lung cancer, and patients with elevated TKT mRNA levels have decreased survival rates compared with those with lower levels157. The representative TKT inhibitor oxythiamine (OT) has been shown to inhibit tumor proliferation by blocking the transition of thiamine to its active form, which is required for the full activity of TKT157,165,167,168. Like PGM2, TKT was found to be important in uridine utilization as an energy source in the absence of glucose24. These results indicate the critical function of TKT in uridine utilization in glucose-deficient cancer cells and suggest its potential as an anticancer target.
R5P to PRPP by PRPS
PRPP synthetase (PRPS) converts R5P into PRPP, which is utilized in nucleotide synthesis169,170,171. In pyrimidine synthesis, the pyrimidine ring is first synthesized and then added to PRPP170. Purine synthesis occurs entirely in the cytosol and the purine ring is directly built onto PRPP172. Hyperactive nucleotide synthesis in cancer cells is a common characteristic, and cancer cells have higher concentrations of dNTPs and NTPs than nonmalignant proliferating cells do2,71. These findings could lead to the development of anticancer agents that target the enzymes involved in nucleotide synthesis. Notably, the loss of TKT has been reported to suppress glycolysis and the TCA cycle, resulting in a reduction in ATP levels and the inhibition of PRPS173. These findings indicate that targeting TKT could lead to the inhibition of PRPS and nucleotide synthesis.
Uridine contribution to glycosylation and phase II detoxification
Another important facet of uridine may be its role in glycosylation, in which sugar nucleotide molecules are utilized as sugar donors. There are 12 sugar nucleotides used in glycosylation reactions in humans, seven of which are UDP bound, including UDP–glucose (UDP–Glc), UDP–glucuronic acid (UDP–GA) and UDP–N-acetylglucosamine (GlcNAc)174. The formation of UDP sugars and subsequent glycosylations are thus tied to uridine metabolism to form UMP and ultimately uridine triphosphate (UTP), which is then used in the UDP sugar-forming condensation reactions. For example, UDP–glucose pyrophosphorylase (UGP) interconverts UTP and glucose 1-phosphate to form inorganic pyrophosphate (PPi) and UDP–Glc175. UDP–Glc can further be converted to UDP–GA via UDP–glucose 6-dehydrogenase (UGDH)176,177.
Thus, uridine formation or uptake is important for the formation of the UDP sugars required for glycosylation. Studies of in vitro cell models have demonstrated that the addition of uridine to cell culture results in dramatic increases in both UDP and cytidine diphosphate sugars and their corresponding glycosylations178,179. Indeed, systemic uridine administration results in an increase in UDP–Glc and UDP–GlcNAc, as well as GlcNAc-glycosylated proteins36,54. Moreover, mitogen stimulation of quiescent cells results in rapid uptake of uridine, which is incorporated into UDP–GA180. These studies indicate that uridine uptake and/or production can be rate limiting in the formation of UDP sugars and the subsequent glycosylation reactions in in vitro tissue culture and in vivo mouse models. Thus, disrupting the contribution of uridine to UDP sugars may have widespread implications for cancer cells, as the glycosylation of proteins and other biomolecules impacts all aspects of the cancer cell surface, including attachment, invasion, surface receptor activity and even interactions with immune cells174. For example, elevated GlcNAc glycosylation of PD-L1 is utilized by cancer cells in immune evasion181; thus, uridine metabolism may contribute to such processes.
Furthermore, the proper glycosylation of proteins is critical for preventing their misfolding and misprocessing; thus, any disruption of the process may induce or sensitize cancer cells to stress on the endoplasmic reticulum (ER) or dysfunction of the Golgi, key sites of glycosylation. In support of the relationship between uridine and ER stress, the unfolded protein response can trigger uridine biosynthesis, whereas disruptions in the uridine–UDP sugar axis can trigger ER stress or Golgi dysfunction177,182,183.
Among the UDP sugars impacted by uridine metabolism, UDP–GA may be particularly relevant to cancer therapy. The formation of UDP–GA is increased in cancer cells177,184, which may be beneficial for tumors via multiple mechanisms, including extracellular matrix modulation, epithelial‒mesenchymal transition and chemoresistance177,185,186. UDP–GA is processed into UDP–xylose by the enzyme UDP–xylose synthase (UXS1), and when UXS1 is disrupted, UDP–GA selectively accumulates in such cancer cells, poisoning them and disrupting their Golgi structure and function177. The notion that excess UDP sugars exert toxic effects suggests that excess production of toxic byproducts may be relevant to the toxicity observed in the presence of excess uridine10,11 and that this could be exploited to exert toxicity in cancer cells.
Uridine metabolism may also be linked to chemoresistance by virtue of its role in UDP–GA production. UDP–GA is the substrate used in a process called ‘glucuronidation’, which is carried out by UDP–glucuronyltransferases (UGTs), a family of 22 enzymes that constitute a phase II detoxification mechanism. The enzymes have differing substrate specificities and can glucuronidate some normal metabolites in addition to xenobiotics (for example, bilirubin)187. In contrast to phase I detoxification (modification), which occurs primarily in the liver, phase II detoxification activity is spread across multiple organs and in cancer cells themselves188. In this reaction, glucuronic acid is conjugated by UGTs to xenobiotic metabolites, which are selectively recognized by the different UGTs. This conjugation has the effects of neutralizing reactive electrophiles and thus detoxifying the compound, as well as rendering them polar and conducive to transport outside of the cell. As such, phase II detoxification represents a mechanism that can increase the chemoresistance of cancer cells. Indeed, various chemotherapeutics have been shown to be glucuronidated and various individual UGTs have been shown to be upregulated in specific tumors. For example, UGT1 and UGT2 isoforms, such as UGT1A1, UGT1A6 and UGT2B17, have been shown to be overexpressed or induced by chemotherapeutics in tumors and can reduce the activity/levels of various chemotherapeutic agents, including irinotecan, sorafenib, raloxifene and tamoxifen188. Additionally, targeting individual UGTs has been shown to induce chemosensitivity in cancer cells188,189.
Uridine has long been indicated as a ‘rescue agent’ against nucleoside chemotherapeutics because of its competitive effect on incorporation into RNA40. The link between uridine metabolism and the sugar nucleotide UDP–GA opens a new line of thinking and direction for future research into how uridine metabolism may regulate the chemoresistance of cancer cells via UDP–GA and glucuronidation; thus, targeting uridine or the various enzymatic steps between uridine and UDP–GA formation could be explored as chemosensitization approaches (Fig. 8).
Uridine is ultimately metabolized to UTP to form UDP–glucose, which in turn can be converted to other sugars, including the glucuronidation substrate UDP–GA. Figure created using Biorender (https://biorender.com/).
Conclusions
Uridine is not only an organism-level ‘currency metabolite’ that is readily available to cells via the plasma4,190, but also a ‘hub molecule’ inside the cell that intersects with numerous intracellular processes24,36,178,179. Uridine can further be obtained via the breakdown/salvage of RNA108,123,191 or via pyrimidine biosynthesis69,192, both of which are highly active processes in cancer cells69. Conversely, uridine itself is a necessary precursor to many biological processes that are central to cancer cell biology. Uridine-derived ribose enables cancer cells to generate energy under conditions of glucose deficiency, contributing to their survival and proliferation24,25. Uridine is rate limiting in the formation of sugar nucleotides that are central to glycosylation reactions54,178,180 and may even be directly linked to chemoresistance because of its metabolic role in the formation of the key phase II detoxification molecule UDP–GA177,186,188. The various enzymes involved in RNA breakdown/salvage, or pyrimidine biosynthesis in many cases, have long been explored for various reasons as cancer therapeutics71,75, but their roles specific to uridine metabolism have not previously been investigated and represent new opportunities to better understand the cancer-related contexts in which they may be most useful. In particular, the recently demonstrated role of RNA polymers as cellular sources of uridine24 indicates that further research is needed into RNA metabolism processes as upstream precursors to uridine metabolism. The enzymes that mediate the processes downstream of uridine leading to glycolysis or UDP sugars24,25,177 have only recently been identified and are also excellent targets for disrupting processes that are integral to cancer cell function.
References
Karle, J. M., Anderson, L. W., Dietrick, D. D. & Cysyk, R. L. Determination of serum and plasma uridine levels in mice, rats, and humans by high-pressure liquid chromatography. Anal. Biochem. 109, 41–46 (1980).
Traut, T. W. Physiological concentrations of purines and pyrimidines. Mol. Cell. Biochem. 140, 1–22 (1994).
van Groeningen, C. J., Peters, G. J., Nadal, J. C., Laurensse, E. & Pinedo, H. M. Clinical and pharmacologic study of orally administered uridine. J. Natl Cancer Inst. 83, 437–441 (1991).
Deng, Y. et al. An adipo–biliary–uridine axis that regulates energy homeostasis. Science 355, eaaf5375 (2017).
Nakagawara, K., Takeuchi, C. & Ishige, K. 5′-CMP and 5′-UMP promote myogenic differentiation and mitochondrial biogenesis by activating myogenin and PGC-1α in a mouse myoblast C2C12 cell line. Biochem. Biophys. Rep. 31, 101309 (2022).
Dixon, T. F. & Purdom, M. Serum 5-nucleotidase. J.Clin. Pathol. 7, 341–343 (1954).
Niedzwicki, J. G., Chu, S. H., el Kouni, M. H., Rowe, E. C. & Cha, S. 5-Benzylacyclouridine and 5-benzyloxybenzylacyclouridine, potent inhibitors of uridine phosphorylase. Biochem. Pharmacol. 31, 1857–1861 (1982).
Choi, J. W. et al. Uridine protects cortical neurons from glucose deprivation-induced death: possible role of uridine phosphorylase. J. Neurotrauma 25, 695–707 (2008).
Jiang, N. & Zhao, Z. Intestinal aging is alleviated by uridine via regulating inflammation and oxidative stress in vivo and in vitro. Cell Cycle 21, 1519–1531 (2022).
Cao, Z. et al. Uridine homeostatic disorder leads to DNA damage and tumorigenesis. Cancer Lett. 372, 219–225 (2016).
Zi, L. et al. Uridine inhibits hepatocellular carcinoma cell development by inducing ferroptosis. J. Clin. Med. 12, 3552 (2023).
Gillies, R. J., Robey, I. & Gatenby, R. A. Causes and consequences of increased glucose metabolism of cancers. J. Nucl. Med. 49, 24s–42s (2008).
Dilliraj, L. N. et al. The evolution of ketosis: potential impact on clinical conditions. Nutrients 14 (2022).
Wahren, J., Felig, P., Cerasi, E. & Luft, R. Splanchnic and peripheral glucose and amino acid metabolism in diabetes mellitus. J. Clin. Invest. 51, 1870–1878 (1972).
Williamson, D. H., Bates, M. W., Page, M. A. & Krebs, H. A. Activities of enzymes involved in acetoacetate utilization in adult mammalian tissues. Biochem. J. 121, 41–47 (1971).
Yen, T. T., Lowry, L., Steinmetz, J. & Wolff, G. L. Physiologic and genetic influences on regulation of glucose metabolism in adipose tissue of mice. Horm. Metab. Res. 2, 161–165 (1970).
Yamamoto, T. et al. Biochemistry of uridine in plasma. Clin. Chim. Acta 412, 1712–1724 (2011).
Gasser, T., Moyer, J. D. & Handschumacher, R. E. Novel single-pass exchange of circulating uridine in rat liver. Science 213, 777–778 (1981).
Yamamoto, T. et al. Effect of ethanol and fructose on plasma uridine and purine bases. Metabolism 46, 544–547 (1997).
Yamamoto, T. et al. Effect of muscular exercise on the concentration of uridine and purine bases in plasma—adenosine triphosphate consumption-induced pyrimidine degradation. Metabolism 46, 1339–1342 (1997).
Yamamoto, T. et al. Effect of beer on the plasma concentrations of uridine and purine bases. Metabolism 51, 1317–1323 (2002).
Baba, S., Yumoto, R., Kawami, M. & Takano, M. Functional expression of equilibrative and concentrative nucleoside transporters in alveolar epithelial cells. Pharmazie 76, 416–421 (2021).
Darnowski, J. W. & Handschumacher, R. E. Tissue uridine pools: evidence in vivo of a concentrative mechanism for uridine uptake. Cancer Res. 46, 3490–3494 (1986).
Skinner, O. S. et al. Salvage of ribose from uridine or RNA supports glycolysis in nutrient-limited conditions. Nat. Metab. 5, 765–776 (2023).
Nwosu, Z. C. et al. Uridine-derived ribose fuels glucose-restricted pancreatic cancer. Nature 618, 151–158 (2023).
Valentin, I., Philippe, M., Lhuguenot, J. & Chagnon, M. Uridine uptake inhibition as a cytotoxicity test for a human hepatoma cell line (HepG2 cells): comparison with the neutral red assay. Toxicology 158, 127–139 (2001).
Hay, N. Reprogramming glucose metabolism in cancer: can it be exploited for cancer therapy? Nat. Rev. Cancer 16, 635–649 (2016).
Christofk, H. R. et al. The M2 splice isoform of pyruvate kinase is important for cancer metabolism and tumour growth. Nature 452, 230–233 (2008).
Noguchi, S. et al. Fluorouracil uptake in triple-negative breast cancer cells: negligible contribution of equilibrative nucleoside transporters 1 and 2. Biopharm. Drug Dispos. 42, 85–93 (2021).
Hioki, M. et al. Contribution of equilibrative nucleoside transporters 1 and 2 to gemcitabine uptake in pancreatic cancer cells. Biopharm. Drug Dispos. 39, 256–264 (2018).
Zhang, J. et al. The role of nucleoside transporters in cancer chemotherapy with nucleoside drugs. Cancer Metastasis Rev. 26, 85–110 (2007).
Santini, D. et al. Human equilibrative nucleoside transporter 1 (hENT1) levels predict response to gemcitabine in patients with biliary tract cancer (BTC). Curr. Cancer Drug Targets 11, 123–129 (2011).
Damaraju, V. L., Sawyer, M. B., Mackey, J. R., Young, J. D. & Cass, C. E. Human nucleoside transporters: biomarkers for response to nucleoside drugs. Nucleosides Nucleotides Nucleic Acids 28, 450–463 (2009).
Galmarini, C. M. et al. In vivo mechanisms of resistance to cytarabine in acute myeloid leukaemia. Br. J. Haematol. 117, 860–868 (2002).
Abraham, A. et al. RNA expression of genes involved in cytarabine metabolism and transport predicts cytarabine response in acute myeloid leukemia. Pharmacogenomics 16, 877–890 (2015).
Yang, Y., Ye, Y., Deng, Y. & Gao, L. Uridine and its role in metabolic diseases, tumors, and neurodegenerative diseases. Front. Physiol. 15, 1360891 (2024).
Pooler, A. M., Guez, D. H., Benedictus, R. & Wurtman, R. J. Uridine enhances neurite outgrowth in nerve growth factor-differentiated PC12 (corrected). Neuroscience 134, 207–214 (2005).
Wiedon, A. et al. Uridine adenosine tetraphosphate (Up4A) is a strong inductor of smooth muscle cell migration via activation of the P2Y2 receptor and cross-communication to the PDGF receptor. Biochem. Biophys. Res. Commun. 417, 1035–1040 (2012).
Pizzorno, G. et al. Homeostatic control of uridine and the role of uridine phosphorylase: a biological and clinical update. Biochim. Biophys. Acta 1587, 133–144 (2002).
Houghton, J. A., Houghton, P. J. & Wooten, R. S. Mechanism of induction of gastrointestinal toxicity in the mouse by 5-fluorouracil, 5-fluorouridine, and 5-fluoro-2′-deoxyuridine. Cancer Res. 39, 2406–2413 (1979).
Liu, Z. et al. Cross-species metabolomic analysis identifies uridine as a potent regeneration promoting factor. Cell Discov. 8, 6 (2022).
Al Absi, H. S., Sacharow, S., Al Zein, N., Al Shamsi, A. & Al Teneiji, A. Hereditary orotic aciduria (HOA): a novel uridine-5-monophosphate synthase (UMPS) mutation. Mol. Genet. Metab. Rep. 26, 100703 (2021).
Frederick, A. et al. Triacetyluridine treats epileptic encephalopathy from CAD mutations: a case report and review. Ann. Clin. Transl. Neurol. 8, 284–287 (2021).
Adant, I. et al. Pyruvate and uridine rescue the metabolic profile of OXPHOS dysfunction. Mol. Metab. 63, 101537 (2022).
Fernández-Moreno, M. et al. Generating Rho-0 cells using mesenchymal stem cell lines. PLoS ONE 11, e0164199 (2016).
King, M. P. & Attardi, G. Human cells lacking mtDNA: repopulation with exogenous mitochondria by complementation. Science 246, 500–503 (1989).
Shoaf, W. T. & Jones, M. E. Uridylic acid synthesis in Ehrlich ascites carcinoma. Properties, subcellular distribution, and nature of enzyme complexes of the six biosynthetic enzymes. Biochemistry 12, 4039–4051 (1973).
Jones, M. E. Pyrimidine nucleotide biosynthesis in animals: genes, enzymes, and regulation of UMP biosynthesis. Annu. Rev. Biochem. 49, 253–279 (1980).
Van Rompay, A. R., Norda, A., Lindén, K., Johansson, M. & Karlsson, A. Phosphorylation of uridine and cytidine nucleoside analogs by two human uridine-cytidine kinases. Mol. Pharmacol. 59, 1181–1186 (2001).
Koizumi, K. et al. Cloning and expression of uridine/cytidine kinase cDNA from human fibrosarcoma cells. Int. J. Mol. Med. 8, 273–278 (2001).
Fukushima, H. et al. 3′-Ethynylcytidine, an RNA polymerase inhibitor, combined with cisplatin exhibits a potent synergistic growth-inhibitory effect via Vaults dysfunction. BMC Cancer 14, 1–12 (2014).
van Kuilenburg, A. B. & Meinsma, R. The pivotal role of uridine–cytidine kinases in pyrimidine metabolism and activation of cytotoxic nucleoside analogues in neuroblastoma. Biochim. Biophys. Acta 1862, 1504–1512 (2016).
Li, Y. et al. Integrative analyses of pyrimidine salvage pathway-related genes revealing the associations between UPP1 and tumor microenvironment. J. Inflamm. Res. 17, 101–119 (2024).
Hawkins, M., Angelov, I., Liu, R., Barzilai, N. & Rossetti, L. The tissue concentration of UDP-N-acetylglucosamine modulates the stimulatory effect of insulin on skeletal muscle glucose uptake. J. Biol. Chem. 272, 4889–4895 (1997).
Shimamoto, Y. et al. Sensitivity of human cancer cells to the new anticancer ribonucleoside TAS–106 is correlated with expression of uridine–cytidine kinase 2. Jpn J. Cancer Res. 93, 825–833 (2002).
Peters, G. J. et al. Metabolism, mechanism of action and sensitivity profile of fluorocyclopentenylcytosine (RX-3117; TV-1360). Invest. New Drugs 31, 1444–1457 (2013).
El Hassouni, B. et al. Uridine cytidine kinase 2 as a potential biomarker for treatment with RX-3117 in pancreatic cancer. Anticancer Res. 39, 3609–3614 (2019).
Malami, I. & Abdul, A. B. Involvement of the uridine cytidine kinase 2 enzyme in cancer cell death: a molecular crosstalk between the enzyme and cellular apoptosis induction. Biomed. Pharmacother. 109, 1506–1510 (2019).
Balboni, B. et al. RX-3117 (fluorocyclopentenyl cytosine): a novel specific antimetabolite for selective cancer treatment. Expert Opin. Investig. Drugs 28, 311–322 (2019).
Choi, W. J. et al. Fluorocyclopentenyl–cytosine with broad spectrum and potent antitumor activity. J. Med. Chem. 55, 4521–4525 (2012).
Sarkisjan, D. et al. The cytidine analog fluorocyclopentenylcytosine (RX-3117) is activated by uridine–cytidine kinase 2. PLoS ONE 11, e0162901 (2016).
Cai, J. et al. Non-metabolic role of UCK2 links EGFR–AKT pathway activation to metastasis enhancement in hepatocellular carcinoma. Oncogenesis 9, 103 (2020).
Fu, Y. et al. The metabolic and non-metabolic roles of UCK2 in tumor progression. Front. Oncol. 12, 904887 (2022).
Coleman, P. F., Suttle, D. P. & Stark, G. R. Purification from hamster cells of the multifunctional protein that initiates de novo synthesis of pyrimidine nucleotides. J. Biol. Chem. 252, 6379–6385 (1977).
Mori, M., Ishida, H. & Tatibana, M. Aggregation states and catalytic properties of the multienzyme complex catalyzing the initial steps of pyrimidine biosynthesis in rat liver. Biochemistry 14, 2622–2630 (1975).
Traut, T. W. & Jones, M. E. Mammalian synthesis of UMP from orotate: the regulation of and conformers of complex U. Adv. Enzyme Regul. 16, 21–41 (1977).
Traut, T. W., Payne, R. C. & Jones, M. E. Dependence of the aggregation and conformation states of uridine 5’-phosphate synthase on pyrimidine nucleotides. Evidence for a regulatory site. Biochemistry 19, 6062–6068 (1980).
Fairbanks, L. D., Bofill, M., Ruckemann, K. & Simmonds, H. A. Importance of ribonucleotide availability to proliferating T-lymphocytes from healthy humans. Disproportionate expansion of pyrimidine pools and contrasting effects of de novo synthesis inhibitors. J. Biol. Chem. 270, 29682–29689 (1995).
Yang, C. et al. De novo pyrimidine biosynthetic complexes support cancer cell proliferation and ferroptosis defence. Nat. Cell Biol. 25, 836–847 (2023).
Wang, S. et al. Bioinformatics analysis and experimental verification of the cancer-promoting effect of DHODH in clear cell renal cell carcinoma. Sci. Rep. 14, 11985 (2024).
Mullen, N. J. & Singh, P. K. Nucleotide metabolism: a pan-cancer metabolic dependency. Nat. Rev. Cancer 23, 275–294 (2023).
Mao, C. et al. DHODH-mediated ferroptosis defence is a targetable vulnerability in cancer. Nature 593, 586–590 (2021).
Mishima, E. et al. DHODH inhibitors sensitize to ferroptosis by FSP1 inhibition. Nature 619, E9–E18 (2023).
Moyer, J. D., Oliver, J. T. & Handschumacher, R. E. Salvage of circulating pyrimidine nucleosides in the rat. Cancer Res. 41, 3010–3017 (1981).
Neff, T. & Blau, C. A. Forced expression of cytidine deaminase confers resistance to cytosine arabinoside and gemcitabine. Exp. Hematol. 24, 1340–1346 (1996).
Ebrahem, Q., Mahfouz, R. Z., Ng, K. P. & Saunthararajah, Y. High cytidine deaminase expression in the liver provides sanctuary for cancer cells from decitabine treatment effects. Oncotarget 3, 1137–1145 (2012).
Shimotohno, K., Kodama, Y., Hashimoto, J. & Miura, K. I. Importance of 5′-terminal blocking structure to stabilize mRNA in eukaryotic protein synthesis. Proc. Natl Acad. Sci. USA 74, 2734–2738 (1977).
Furuichi, Y., LaFiandra, A. & Shatkin, A. J. 5′-Terminal structure and mRNA stability. Nature 266, 235–239 (1977).
Sachs, A. B. Messenger RNA degradation in eukaryotes. Cell 74, 413–421 (1993).
Bird, J. G. et al. The mechanism of RNA 5′ capping with NAD+, NADH and desphospho-CoA. Nature 535, 444–447 (2016).
Bird, J. G. et al. Highly efficient 5′ capping of mitochondrial RNA with NAD+ and NADH by yeast and human mitochondrial RNA polymerase. eLife 7, e42179 (2018).
Decker, C. J. & Parker, R. A turnover pathway for both stable and unstable mRNAs in yeast: evidence for a requirement for deadenylation. Genes Dev. 7, 1632–1643 (1993).
Popp, M. W. & Maquat, L. E. Organizing principles of mammalian nonsense-mediated mRNA decay. Annu. Rev. Genet. 47, 139–165 (2013).
Kraft, C., Deplazes, A., Sohrmann, M. & Peter, M. Mature ribosomes are selectively degraded upon starvation by an autophagy pathway requiring the Ubp3p/Bre5p ubiquitin protease. Nat. Cell Biol. 10, 602–610 (2008).
Sander, G., Topp, H., Heller-Schöch, G., Wieland, J. & Schöch, G. Ribonucleic acid turnover in man:RNA catabolites in urine as measure for the metabolism of each of the three major species of RNA. Clin. Sci. 71, 367–374 (1986).
Kaberdin, V. R., Singh, D. & Lin-Chao, S. Composition and conservation of the mRNA-degrading machinery in bacteria. J. Biomed. Sci. 18, 23 (2011).
MacIntosh, G. C. & Bassham, D. C. The connection between ribophagy, autophagy and ribosomal RNA decay. Autophagy 7, 662–663 (2011).
Mugridge, J. S., Coller, J. & Gross, J. D. Structural and molecular mechanisms for the control of eukaryotic 5′–3′ mRNA decay. Nat. Struct. Mol. Biol. 25, 1077–1085 (2018).
Yamashita, A. et al. Concerted action of poly(A) nucleases and decapping enzyme in mammalian mRNA turnover. Nat. Struct. Mol. Biol. 12, 1054–1063 (2005).
Collart, M. A. & Panasenko, O. O. The Ccr4–Not complex. Gene 492, 42–53 (2012).
Sohn, E. J. et al. CNOT2 promotes proliferation and angiogenesis via VEGF signaling in MDA-MB-231 breast cancer cells. Cancer Lett. 412, 88–98 (2018).
Jung, J. H., Lee, D., Ko, H. M. & Jang, H. J. Inhibition of CNOT2 induces apoptosis via MID1IP1 in colorectal cancer cells by activating p53. Biomolecules 11, 1492 (2021).
Kim, E. O., Kang, S. E., Choi, M., Rhee, K. J. & Yun, M. CCR4–NOT transcription complex subunit 2 regulates TRAIL sensitivity in non‑small‑cell lung cancer cells via the STAT3 pathway. Int. J. Mol. Med. 45, 324–332 (2020).
Ghashghaei, M. et al. Translation efficiency driven by CNOT3 subunit of the CCR4–NOT complex promotes leukemogenesis. Nat. Commun. 15, 2340 (2024).
Jing, L. et al. CNOT3 contributes to cisplatin resistance in lung cancer through inhibiting RIPK3 expression. Apoptosis 24, 673–685 (2019).
Jing, L. et al. Targeting the up-regulated CNOT3 reverses therapeutic resistance and metastatic progression of EGFR-mutant non-small cell lung cancer. Cell Death Discov. 9, 406 (2023).
Faraji, F. et al. Post-transcriptional control of tumor cell autonomous metastatic potential by CCR4–NOT deadenylase CNOT7. PLoS Genet. 12, e1005820 (2016).
Song, M.-G., Li, Y. & Kiledjian, M. Multiple mRNA decapping enzymes in mammalian cells. Mol. Cell 40, 423–432 (2010).
Garneau, N. L., Wilusz, J. & Wilusz, C. J. The highways and byways of mRNA decay. Nat. Rev. Mol. Cell Biol. 8, 113–126 (2007).
Wang, Z. & Kiledjian, M. Functional link between the mammalian exosome and mRNA decapping. Cell 107, 751–762 (2001).
Liu, H., Rodgers, N. D., Jiao, X. & Kiledjian, M. The scavenger mRNA decapping enzyme DcpS is a member of the HIT family of pyrophosphatases. EMBO J. 21, 4699–4708 (2002).
Milac, A. L., Bojarska, E. & Wypijewska del Nogal, A. Decapping scavenger (DcpS) enzyme: advances in its structure, activity and roles in the cap-dependent mRNA metabolism. Biochim. Biophys. Acta 1839, 452–462 (2014).
Gu, M. et al. Insights into the structure, mechanism, and regulation of scavenger mRNA decapping activity. Mol. Cell 14, 67–80 (2004).
Chen, N., Walsh, M. A., Liu, Y., Parker, R. & Song, H. Crystal structures of human DcpS in ligand-free and m7GDP-bound forms suggest a dynamic mechanism for scavenger mRNA decapping. J. Mol. Biol. 347, 707–718 (2005).
Yamauchi, T. et al. Genome-wide CRISPR–Cas9 screen identifies leukemia-specific dependence on a pre-mRNA metabolic pathway regulated by DCPS. Cancer Cell 33, 386–400.e5 (2018).
Shalem, O. et al. Genome-scale CRISPR–Cas9 knockout screening in human cells. Science 343, 84–87 (2014).
Swartzel, J. C. et al. Targeted degradation of mRNA decapping enzyme DcpS by a VHL-recruiting PROTAC. ACS Chem. Biol. 17, 1789–1798 (2022).
Li, Y. & Kiledjian, M. Regulation of mRNA decapping. Wiley Interdiscip. Rev. 1, 253–265 (2010).
Borbolis, F. & Syntichaki, P. Biological implications of decapping: beyond bulk mRNA decay. FEBS J. 289, 1457–1475 (2022).
van Dijk, E. et al. Human Dcp2: a catalytically active mRNA decapping enzyme located in specific cytoplasmic structures. EMBO J. 21, 6915–6924 (2002).
Wang, Z., Jiao, X., Carr-Schmid, A. & Kiledjian, M. The hDcp2 protein is a mammalian mRNA decapping enzyme. Proc. Natl Acad. Sci. USA 99, 12663–12668 (2002).
Nagarajan, V. K., Jones, C. I., Newbury, S. F. & Green, P. J. XRN 5′→3′ exoribonucleases: structure, mechanisms and functions. Biochim. Biophys. Acta 1829, 590–603 (2013).
Hsu, C. L. & Stevens, A. Yeast cells lacking 5′->3′ exoribonuclease 1 contain mRNA species that are poly(A) deficient and partially lack the 5′ cap structure. Mol. Cell. Biol. 13, 4826–4835 (1993).
Zou, T. et al. XRN1 deletion induces PKR-dependent cell lethality in interferon-activated cancer cells. Cell Rep. 43, 113600 (2024).
Ran, X. B. et al. Targeting RNA exonuclease XRN1 potentiates efficacy of cancer immunotherapy. Cancer Res. 83, 922–938 (2023).
Wu, H. L. et al. Targeting nucleotide metabolism: a promising approach to enhance cancer immunotherapy. J. Hematol. Oncol. 15, 45 (2022).
Silverman, R. H. Viral encounters with 2′,5′-oligoadenylate synthetase and RNase L during the interferon antiviral response. J. Virol. 81, 12720–12729 (2007).
Anderson, B. R. et al. Nucleoside modifications in RNA limit activation of 2′-5′-oligoadenylate synthetase and increase resistance to cleavage by RNase L. Nucleic Acids Res. 39, 9329–9338 (2011).
Hovanessian, A. G., Brown, R. E. & Kerr, I. M. Synthesis of low molecular weight inhibitor of protein synthesis with enzyme from interferon-treated cells. Nature 268, 537–540 (1977).
Kerr, I. M. & Brown, R. E. pppA2′p5′A2′p5′A: an inhibitor of protein synthesis synthesized with an enzyme fraction from interferon-treated cells. Proc. Natl Acad. Sci. USA 75, 256–260 (1978).
Lin, R. J. et al. Distinct antiviral roles for human 2′,5′-oligoadenylate synthetase family members against dengue virus infection. J. Immunol. 183, 8035–8043 (2009).
Rutherford, M. N., Hannigan, G. E. & Williams, B. R. Interferon-induced binding of nuclear factors to promoter elements of the 2–5A synthetase gene. EMBO J. 7, 751–759 (1988).
Banerjee, S. et al. RNase L is a negative regulator of cell migration. Oncotarget 6, 44360 (2015).
Molinaro, R. J. et al. Selection and cloning of poly (rC)-binding protein 2 and Raf kinase inhibitor protein RNA activators of 2′,5′-oligoadenylate synthetase from prostate cancer cells. Nucleic Acids Res. 34, 6684–6695 (2006).
Tanaka, N. et al. Structural basis for recognition of 2′,5′-linked oligoadenylates by human ribonuclease L. EMBO J. 23, 3929–3938 (2004).
Zhang, A. et al. RNase L restricts the mobility of engineered retrotransposons in cultured human cells. Nucleic Acids Res. 42, 3803–3820 (2014).
Zhou, A. et al. Interferon action and apoptosis are defective in mice devoid of 2′,5′-oligoadenylate-dependent RNase L. EMBO J. 16, 6355–6363 (1997).
Chakrabarti, A. et al. RNase L activates the NLRP3 inflammasome during viral infections. Cell Host Microbe 17, 466–477 (2015).
Chakrabarti, A., Ghosh, P. K., Banerjee, S., Gaughan, C. & Silverman, R. H. RNase L triggers autophagy in response to viral infections. J. Virol. 86, 11311–11321 (2012).
Dayal, S. et al. RNase L suppresses androgen receptor signaling, cell migration and matrix metalloproteinase activity in prostate cancer cells. Int. J. Mol. Sci. 18, 529 (2017).
Yin, H., Jiang, Z., Wang, S. & Zhang, P. IFN-γ restores the impaired function of RNase L and induces mitochondria-mediated apoptosis in lung cancer. Cell Death Dis. 10, 642 (2019).
Evans, D. R. & Guy, H. I. Mammalian pyrimidine biosynthesis: fresh insights into an ancient pathway. J. Biol. Chem. 279, 33035–33038 (2004).
Walter, M. & Herr, P. Re-discovery of pyrimidine salvage as target in cancer therapy. Cells 11, 739 (2022).
Brandão, T. A. & Richard, J. P. Orotidine 5′-monophosphate decarboxylase: the operation of active site chains within and across protein subunits. Biochemistry 59, 2032–2040 (2020).
Zhang, Y. et al. Targeting glucose metabolism enzymes in cancer treatment: current and emerging strategies. Cancers 14, 4568 (2022).
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).
Kelloff, G. J. et al. Progress and promise of FDG–PET imaging for cancer patient management and oncologic drug development. Clin. Cancer Res. 11, 2785–2808 (2005).
Luo, Y. et al. A nanounit strategy disrupts energy metabolism and alleviates immunosuppression for cancer therapy. Nano Lett. 22, 6418–6427 (2022).
Yu, J. et al. Advanced cancer starvation therapy by simultaneous deprivation of lactate and glucose using a MOF nanoplatform. Adv. Sci. 8, 2101467 (2021).
Zhang, J. et al. TME-triggered MnSiO3@ Met@ GOx nanosystem for ATP dual-inhibited starvation/chemodynamic synergistic therapy. Biomaterials 287, 121682 (2022).
Keenan, M. M. & Chi, J.-T. Alternative fuels for cancer cells. Cancer J. 21, 49–55 (2015).
Roosild, T. P., Castronovo, S., Villoso, A., Ziemba, A. & Pizzorno, G. A novel structural mechanism for redox regulation of uridine phosphorylase 2 activity. J. Struct. Biol. 176, 229–237 (2011).
Zhao, Y. et al. 5-Fluorouracil enhances the antitumor activity of the glutaminase inhibitor CB-839 against PIK3CA-mutant colorectal cancers. Cancer Res. 80, 4815–4827 (2020).
Yan, R., Wan, L., Pizzorno, G. & Cao, D. Uridine phosphorylase in breast cancer: a new prognostic factor. Front. Biosci. 11, 2759–2766 (2006).
Wang, X. et al. UPP1 promotes lung adenocarcinoma progression through epigenetic regulation of glycolysis. Aging Dis. 13, 1488 (2022).
Miyashita, H. et al. Uridine phosphorylase is a potential prognostic factor in patients with oral squamous cell carcinoma. Cancer 94, 2959–2966 (2002).
Guan, Y., Bhandari, A., Zhang, X. & Wang, O. Uridine phosphorylase 1 associates to biological and clinical significance in thyroid carcinoma cell lines. J. Cell. Mol. Med. 23, 7438–7448 (2019).
da Silva, E. F. G. et al. CPBMF65, a synthetic human uridine phosphorylase-1 inhibitor, reduces HepG2 cell proliferation through cell cycle arrest and senescence. Invest. New Drugs 38, 1653–1663 (2020).
Choi, J. W. et al. Uridine prevents the glucose deprivation-induced death of immunostimulated astrocytes via the action of uridine phosphorylase. Neurosci. Res. 56, 111–118 (2006).
Du, W. et al. UPP1 enhances bladder cancer progression and gemcitabine resistance through AKT. Int. J. Biol. Sci. 20, 1389–1409 (2024).
Zhang, Y. et al. Identification of a liver-specific uridine phosphorylase that is regulated by multiple lipid-sensing nuclear receptors. Mol. Endocrinol. 18, 851–862 (2004).
Kong, X. et al. Peroxisome proliferator-activated receptor gamma coactivator-1 alpha enhances antiproliferative activity of 5′-deoxy-5-fluorouridine in cancer cells through induction of uridine phosphorylase. Mol. Pharmacol. 76, 854–860 (2009).
Hu, Y. et al. Ganoderma lucidum phosphoglucomutase is required for hyphal growth, polysaccharide production, and cell wall integrity. Appl. Microbiol. Biotechnol. 102, 1911–1922 (2018).
Zheng, Z. et al. Meta-analysis of the effect of PGM on survival prognosis of tumor patients. Front. Oncol. 12, 1060372 (2022).
Kvam, C., Olsvik, E. S., Mckinley-Mckee, J. & Saether, O. Studies on recombinant Acetobacter xylinum α-phosphoglucomutase. Biochem. J. 326, 197–203 (1997).
Yu, H., Yu, Z., Qin, C., Bian, T. & Shi, M. High expression of glycolysis-related PGM2 gene in relation to poor prognosis and deficient immune cells infiltration in lung adenocarcinoma: a study based on bioinformatics analysis. J. Thoracic Dis. 14, 3488 (2022).
Hao, S. et al. The role of transketolase in human cancer progression and therapy. Biomed. Pharmacother. 154, 113607 (2022).
Qin, Z. et al. Transketolase (TKT) activity and nuclear localization promote hepatocellular carcinoma in a metabolic and a non-metabolic manner. J. Exp. Clin. Cancer Res. 38, 1–21 (2019).
McCommis, K. S. & Finck, B. N. Mitochondrial pyruvate transport: a historical perspective and future research directions. Biochem. J. 466, 443–454 (2015).
Pelicano, H., Martin, D., Xu, R. & Huang, P. Glycolysis inhibition for anticancer treatment. Oncogene 25, 4633–4646 (2006).
Nie, H. et al. The short isoform of PRLR suppresses the pentose phosphate pathway and nucleotide synthesis through the NEK9–Hippo axis in pancreatic cancer. Theranostics 11, 3898 (2021).
Shukla, S. K. et al. MUC1 and HIF-1alpha signaling crosstalk induces anabolic glucose metabolism to impart gemcitabine resistance to pancreatic cancer. Cancer Cell 32, 71–87.e7 (2017).
Samatiwat, P., Prawan, A., Senggunprai, L., Kukongviriyapan, U. & Kukongviriyapan, V. Nrf2 inhibition sensitizes cholangiocarcinoma cells to cytotoxic and antiproliferative activities of chemotherapeutic agents. Tumor Biol. 37, 11495–11507 (2016).
Yang, H. et al. MicroRNA-497 regulates cisplatin chemosensitivity of cervical cancer by targeting transketolase. Am. J. Cancer Res. 6, 2690 (2016).
Dasgupta, S. et al. Metabolic enzyme PFKFB4 activates transcriptional coactivator SRC-3 to drive breast cancer. Nature 556, 249–254 (2018).
Zhao, F. et al. Imatinib resistance associated with BCR-ABL upregulation is dependent on HIF-1α-induced metabolic reprograming. Oncogene 29, 2962–2972 (2010).
Raı̈s, B. et al. Oxythiamine and dehydroepiandrosterone induce a G1 phase cycle arrest in Ehrlich’s tumor cells through inhibition of the pentose cycle. FEBS Lett. 456, 113–118 (1999).
Liu, C.-L. et al. Targeting the pentose phosphate pathway increases reactive oxygen species and induces apoptosis in thyroid cancer cells. Mol. Cell. Endocrinol. 499, 110595 (2020).
Hove-Jensen, B. et al. Phosphoribosyl diphosphate (PRPP): biosynthesis, enzymology, utilization, and metabolic significance. Microbiol. Mol. Biol. Rev. 81, 00040–16 (2017).
Kornberg, A., Lieberman, I. & Simms, E. S. Enzymatic synthesis and properties of 5-phosphoribosylpyrophosphate. J. Biol. Chem. 215, 389–402 (1955).
Khorana, H. G., Fernandes, J. F. & Kornberg, A. Pyrophosphorylation of ribose 5-phosphate in the enzymatic synthesis of 5-phosphorylribose 1-pyrophosphate. J. Biol. Chem. 230, 941–948 (1958).
Villa, E., Ali, E., Sahu, U. & Ben-Sahra, I. Cancer cells tune the signaling pathways to empower de novo synthesis of nucleotides. Cancers 10.3390/cancers11050688 (2019).
Guo, X. et al. Metabolic plasticity, essentiality and therapeutic potential of ribose-5-phosphate synthesis in Toxoplasma gondii. Nat. Commun. 15, 2999 (2024).
Reily, C., Stewart, T. J., Renfrow, M. B. & Novak, J. Glycosylation in health and disease. Nat. Rev. Nephrol. 15, 346–366 (2019).
Führing, J. I. et al. A quaternary mechanism enables the complex biological functions of octameric human UDP–glucose pyrophosphorylase, a key enzyme in cell metabolism. Sci. Rep. 5, 9618 (2015).
Spicer, A. P., Kaback, L. A., Smith, T. J. & Seldin, M. F. Molecular cloning and characterization of the human and mouse UDP−glucose dehydrogenase genes. J. Biol. Chem. 273, 25117–25124 (1998).
Doshi, M. B. et al. Disruption of sugar nucleotide clearance is a therapeutic vulnerability of cancer cells. Nature 623, 625–632 (2023).
Wong, N. S. et al. An investigation of intracellular glycosylation activities in CHO cells: effects of nucleotide sugar precursor feeding. Biotechnol. Bioeng. 107, 321–336 (2010).
Gramer, M. J. et al. Modulation of antibody galactosylation through feeding of uridine, manganese chloride, and galactose. Biotechnol. Bioeng. 108, 1591–1602 (2011).
Zaharevitz, D. W., Chisena, C. A. & Cysyk, R. L. Rapid increase of cellular UDP–glucuronide after mitogen stimulation of quiescent 3T3 mouse fibroblasts. Biochem. Int. 20, 1067–1076 (1990).
Zhu, Q. et al. O-GlcNAcylation promotes tumor immune evasion by inhibiting PD-L1 lysosomal degradation. Proc. Natl Acad. Sci. USA 120, e2216796120 (2023).
Deng, Y. et al. Adipocyte Xbp1s overexpression drives uridine production and reduces obesity. Mol. Metab. 11, 1–17 (2018).
Flores-Diaz, M. et al. A cellular UDP–glucose deficiency causes overexpression of glucose/oxygen-regulated proteins independent of the endoplasmic reticulum stress elements. J. Biol. Chem. 279, 21724–21731 (2004).
Wang, T. P., Pan, Y. R., Fu, C. Y. & Chang, H. Y. Down-regulation of UDP–glucose dehydrogenase affects glycosaminoglycans synthesis and motility in HCT-8 colorectal carcinoma cells. Exp. Cell Res. 316, 2893–2902 (2010).
Fan, M. et al. UDP–glucose dehydrogenase supports autophagy-deficient PDAC growth via increasing hyaluronic acid biosynthesis. Cell Rep. 43, 113808 (2024).
Harrington, B. S. et al. UGDH promotes tumor-initiating cells and a fibroinflammatory tumor microenvironment in ovarian cancer. J. Exp. Clin. Cancer Res. 42, 270 (2023).
Yang, G. et al. Glucuronidation: driving factors and their impact on glucuronide disposition. Drug Metab. Rev. 49, 105–138 (2017).
Allain, E. P., Rouleau, M., Lévesque, E. & Guillemette, C. Emerging roles for UDP–glucuronosyltransferases in drug resistance and cancer progression. Br. J. Cancer 122, 1277–1287 (2020).
Ascierto, M. L. et al. The intratumoral balance between metabolic and immunologic gene expression is associated with anti-PD-1 response in patients with renal cell carcinoma. Cancer Immunol. Res. 4, 726–733 (2016).
Jankowski, V. et al. Uridine adenosine tetraphosphate: a novel endothelium-derived vasoconstrictive factor. Nat. Med. 11, 223–227 (2005).
Mugridge, J. S., Coller, J. & Gross, J. D. Structural and molecular mechanisms for the control of eukaryotic 5′-3′ mRNA decay. Nat. Struct. Mol. Biol. 25, 1077–1085 (2018).
Zhang, Y., Guo, S., Xie, C. & Fang, J. Uridine metabolism and its role in glucose, lipid, and amino acid homeostasis. Biomed. Res. Int. 2020, 7091718 (2020).
Acknowledgements
We regret that many key findings related to uridine metabolism, glycosylation and RNA decay could not be included in the review owing to space limitations. K.-M.C. and J.-H.Y. are supported by startup funds from the University of Oklahoma. D.K. is supported by the NCI (CA269711) and NIGMS (GM148832).
Author information
Authors and Affiliations
Corresponding authors
Ethics declarations
Competing interests
The authors declare no competing interests.
Additional information
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/.
About this article
Cite this article
Choi, KM., Berard, B.A., Yoon, JH. et al. Uridine as a hub in cancer metabolism and RNA biology. Exp Mol Med 57, 1651–1662 (2025). https://doi.org/10.1038/s12276-025-01402-7
Received:
Revised:
Accepted:
Published:
Version of record:
Issue date:
DOI: https://doi.org/10.1038/s12276-025-01402-7
