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Published in final edited form as: Wiley Interdiscip Rev Syst Biol Med. 2018 Jun 11;10(6):e1426. doi: 10.1002/wsbm.1426

Cell cycle regulation of folate-mediated one-carbon metabolism

Xu Lan 1, Martha S Field 1, Patrick J Stover 1
PMCID: PMC11875019  NIHMSID: NIHMS2057343  PMID: 29889360

Abstract

Folate-mediated one-carbon metabolism (FOCM) comprises a network of interconnected folate-dependent metabolic pathways responsible for serine and glycine interconversion, de novo purine synthesis, de novo thymidylate synthesis and homocysteine remethylation to methionine. These pathways are compartmentalized in the cytosol, nucleus and mitochondria. Individual enzymes within the FOCM network compete for folate cofactors because intracellular folate concentrations are limiting. Although there are feedback mechanisms that regulate the partitioning of folate cofactors among the folate-dependent pathways, less recognized is the impact of cell cycle regulation on FOCM. This review summarizes the evidence for temporal regulation of expression, activity and cellular localization of enzymes and pathways in the FOCM network in mammalian cells through the cell cycle.

This article is categorized under:

Biological Mechanisms > Metabolism

Physiology > Mammalian Physiology in Health and Disease

Keywords: cell cycle, folate, one-carbon metabolism

1 |. INTRODUCTION

Folate-mediated one-carbon metabolism (FOCM) comprises a network of interconnected folate-dependent metabolic pathways responsible for serine and glycine interconversion, de novo purine synthesis, de novo thymidylate (dTMP) synthesis and homocysteine remethylation to methionine. These pathways are compartmentalized in the cytosol, nucleus and mitochondria. They are interconnected through their dependency on one-carbon (1C) units derived primarily from serine and glycine cataboism, and through their reliance on folate cofactors, which carry and chemically activate 1C units at the oxidation states of formate, formaldehyde and methanol (Fox & Stover, 2008). Intracellular folate concentrations are limiting for individual enzymatic reactions within the FOCM network, because the cellular concentration of folate-dependent enzymes exceeds by several fold the concentration of intracellular folate. Hence, the biosynthetic reactions of FOCM compete for folate cofactors (Suh, Oppenheim, Girgis, & Stover, 2000). Cells preferentially direct the flux of 1C units to meet changes in cellular demands due to stress conditions, including nutrient deprivation and disease states such as cancer (Nilsson et al., 2014; Paone et al., 2014). Although there are feedback mechanisms that regulate the partitioning of folate cofactors among the folate-dependent pathways (Field, Szebenyi, & Stover, 2006; Stover & Schirch, 1991), there are also temporal differences in the activity and expression of the enzymes that constitute the various pathways within FOCM. Regulation of the partitioning of folate cofactors among FOCM pathways is essential to address metabolic needs that fluctuate through cell cycle progression, such as increased demand of dTMP for DNA replication during S phase of the cell cycle (Anderson, Woeller, Chiang, Shane, & Stover, 2012). Temporal changes in FOCM metabolic inputs and outputs can be achieved through the regulation of FOCM enzyme levels at the level of transcription (Table 1), translation and post-translational modification as well as regulation of FOCM enzyme subcellular localization. This review synthesizes the evidence for pathway-specific and cell cycle-dependent regulation of enzymes in or related to the FOCM pathway. Specifically, we focus on the temporal changes in the expression and cellular localization of these enzymes in mammalian cells as a function of the cell cycle.

TABLE 1.

Genes involved in one-carbon metabolism network are regulated by transcription factors in a cell cycle-dependent manner

Gene name Transcription factors Cell cycle stage of transcription induction Binding site Reference
CAD Myc/Max G1/S phase Promoter Boyd and Farnham (1997)
DHFR E2F Promoter Abali, Skacel, Celikkaya, and Hsieh (2008)
Sp1 Throughout cell cycle Promoter Good, Dimri, Campisi, and Chen (1996), Slansky and Farnham (1996)
MTHFD1 FOXM1 Promoter Grant et al. (2013)
MTHFS FOXM1 Grant et al. (2013)
RRM1 E2F DeGregori, Kowalik, and Nevins (1995), Ishida et al. (2001)
RRM2 E2F S phase (Chabes, Björklund, & Thelander, 2004) Chabes et al. (2004), DeGregori et al. (1995), Hurford, Cobrinik, Lee, and Dyson (1997), Ishida et al. (2001)
TK1 E2F Ishida et al. (2001)
FOXM1 Grant et al. (2013)
TYMS E2F G1 to S transition (Lee & Johnson, 2000) DeGregori et al. (1995), Ishida et al. (2001), Lee and Johnson (2000), Polager, Kalma, Berkovich, and Ginsberg, (2002)
FOXM1 Grant et al. (2013), Huynh et al. (2011)
LSF G1/S Promoter and intronic regions Powell, Rudge, Zhu, Johnson, and Hansen (2000)
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2 |. FOCM IN THE CYTOSOL

Folate-activated 1C units in the FOCM network can be generated directly by the conversion of tetrahydrofolate (THF) and serine to glycine and 5,10-methylene-THF catalyzed by serine hydroxymethyltransferase (SHMT1 or SHMT2α) in the cytosol and nucleus. Alternatively, 10-formylTHF is formed from THF, ATP and formate catalyzed by methylenetetrahydrofolate dehydrogenase 1 (MTHFD1) in the cytosol and nucleus. Formate is a major source of 1C units, and is derived primarily from THF-dependent serine and glycine catabolism in the mitochondria. Elevated plasma formate concentrations and decreased rates of de novo formate production were observed in the rats fed on folate-deficient diet compared with rats fed on folate-replete diet (Morrow et al., 2015). The relative contributions of endogenous formate production and exogenous formate uptake on cellular formate concentration remain unclear. Other sources of 1C units include the degradation of histidine in the cytosol (Ducker & Rabinowitz, 2017; Solans, Estivill, & de la Luna, 2000), and degradation of choline-derived methyl-glycine species, including betaine, dimethylglycine (DMG), and sarcosine (Figure 1). Choline can be oxidized to betaine via choline dehydrogenase and betaine aldehyde dehydrogenase. In the cytosol, betaine-homocysteine S-methyltransferase provides a 1C unit through the conversion of betaine and homocysteine to DMG and methionine in a pathway that is folate independent. DMG is catabolized to sarcosine and glycine in the mitochondria through dimethylglycine dehydrogenase and sarcosine dehydrogenase respectively, generating additional folate-activated 1C units. In the cytosol, formate-derived, folate-activated 1C units enter into the FOCM network and are incorporated into the end products of de novo purine biosynthesis, de novo dTMP biosynthesis, and the remethylation of homocysteine to methionine. Methionine can be converted to S-adenosyl-methionine, a cofactor that donates methyl groups for numerous methylation reactions, including methylation of DNA, RNA, proteins, phospholipids, and neurotransmitters (Clarke & Banfield, 2001; Selhub, 1999).

FIGURE 1.

FIGURE 1

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Compartmentation of folate-mediated one-carbon metabolism. One-carbon units in FOCM are derived from serine and histidine catabolism in the cytosol, and from the catabolism of serine, glycine and choline-derived methyl-glycine species in the mitochondria. FOCM in the cytoplasm is required for de novo purine synthesis, for de novo thymidylate synthesis and for homocysteine remethylation to methionine. FOCM in the nucleus is required for de novo thymidylate synthesis at sites of DNA replication. The de novo thymidylate synthesis pathway is also present in the mitochondria [Correction added on 25 June 2018, after first online publication: Figure 1 has been updated to include missing labels in some of the gray boxes.]

2.1 |. Methylenetetrahydrofolate dehydrogenase 1

2.1.1 |. MTHFD1 expression

MTHFD1 is a trifunctional enzyme with 5, 10-methylene-THF dehydrogenase, 5,10-methenylTHF cyclohydrolase and 10-formylTHF synthetase activities. In the cytosol, MTHFD1 is the source of formate-derived 1C units which are required for de novo purine synthesis (through 10-formylTHF synthesis) as well as de novo dTMP synthesis and homocysteine remethylation into methionine (through methylene-THF synthesis). The MTHFD1 gene is predominantly expressed in G1/S and G2 phase of the cell cycle in human foreskin fibroblasts (Bar-Joseph et al., 2008). Its promoter is bound by the G2/M transcription factor FOXM1 in U2OS and Hela cells (Grant et al., 2013), although the mechanism of cell cycle regulation of MTHFD1 gene expression remains unclear. Future study is needed to assess the changes in MTHFD1 protein levels throughout the cell cycle.

2.1.2 |. MTHFD1 subcellular localization

MTHFD1 protein translocates into the nucleus during S phase, and the partitioning of MTHFD1 between the cytosol and nucleus is influenced by cellular folate levels, with increased MTHFD1 nuclear localization observed in response to folate depletion (Field et al., 2014; Field, Kamynina, Watkins, Rosenblatt, & Stover, 2015). A human MTHFD1-green fluorescent protein (GFP) fusion protein expressed in HeLa cells was observed in the nucleus, and the levels of MTHFD1-GFP fusion protein in the nucleus were nearly twofold higher in S phase-arrested cells compared to cell arrested in either G1 or G2/M phase. Endogenous MTHFD1 protein nuclear localization also increased during S phase in Hela cells (Field et al., 2014). In mouse liver, MTHFD1 nuclear localization was increased when mice were fed a diet lacking folate (Field et al., 2014). Human fibroblasts expressing an MTHFD1 variant with reduced activity exhibited increased MTHFD1 nuclear localization, suggesting MTHFD1 nuclear localization supports de novo dTMP biosynthesis in the nucleus at the expense of homocysteine remethylation pathway in the cytosol, especially during folate deficiency (Field et al., 2015).

2.2 |. The de novo dTMP synthesis pathway

2.2.1 |. Pyrimidine biosynthesis

Pyrimidine biosynthesis is regulated through the cell cycle to meet increasing cellular demand for both cytosine and thymidine deoxyribonucleotides during DNA replication. The pyrimidine biosynthesis pathway was shown to be upregulated twofold prior to entry into S phase and decreased upon exit from S phase in baby hamster kidney cells (Sigoillot, Berkowski, Sigoillot, Kotsis, & Guy, 2003). Similarly, pyrimidine nucleotide concentrations are increased during growth in rat livers (Jackson, Lui, Boritzki, Morris, & Weber, 1980).

The carbamoyl-phosphate synthetase 2 gene (CAD) encodes a trifunctional protein that catalyzes the first three activities of the six-step pyrimidine biosynthesis pathway: carbamoyl-phosphate synthetase (CPS II), aspartate transcarbamoylase (ATCase), and dihydroorotase. The cad gene is regulated throughout the cell cycle at both the transcriptional and post-transcriptional levels. Steady-state cad RNA levels increased 13-fold, peaking during mid to late G1 phase of the cell cycle, in serum stimulated Syrian hamster ts13 cells (Rao & Davidson, 1988). Newly synthesized CAD protein increased in parallel (Rao & Davidson, 1988). However, the rate of cad transcription increased only about twofold (Rao & Davidson, 1988), indicating that post-transcriptional regulation plays an important role. Similarly, in serum stimulated mouse fibroblasts, steady-state levels of cad mRNA increased 10-fold whereas its transcription increased only threefold (Rao & Church, 1988). When teratocarcinoma cells were induced into differentiation, steady-state levels of cad mRNA decreased by sevenfold while the rate of cad mRNA transcription remained the same, suggesting that expression of the cad gene is cell-growth dependent and regulated at the post-transcriptional level (Rao, Church, & Davidson, 1988). Several transcription factors have been shown to regulate cad gene expression. Binding of Myc/Max at cad promoter is essential for growth-induced cad expression regulation (Boyd & Farnham, 1997). The cad gene is also under the regulation of induction by a nonclassical ERα/Sp1-mediated pathway and repression by HIF-1α(Chen, Lai, Sun, & Tsai, 2005; Khan, Abdelrahim, Samudio, & Safe, 2003).

The activities of enzymes in the de novo pyrimidine synthesis pathway are direct targets of cell cycle regulation. Increased ATCase activity was observed following increases in the cad transcription rate and mRNA level in serum stimulated mouse fibroblasts (Rao & Church, 1988). CPS II, ATCase and orotate phosphoribosyl transferase (OPRTase, another enzyme in pyrimidine biosynthetic pathway) have distinct peaks of enzymatic activity during S phase and diminished activity during the G2/M phase in rat hepatoma cells. More specifically, CPS II and ATCase activities were shown to be increased rapidly during early G1, whereas OPRTase activity did not increase until late G1 (Mitchell & Hoogenraad, 1975).

CAD activity is also regulated throughout the cell cycle by phosphorylation and dephosphorylation signaling cascades. Phosphorylated CAD stimulates de novo pyrimidine synthesis and progression through S phase of the cell cycle in mammalian cells mediated by the activation of mTORC1 (Robitaille et al., 2013). Ribosomal protein s6 kinase 1, a mTORC1 target, phosphorylates S1859 on CAD, promoting CAD oligomerization (Ben-Sahra, Howell, Asara, & Manning, 2013; Robitaille et al., 2013). In baby hamster kidney cells, CAD was activated by MAPK phosphorylation just before the onset of S phase, and then rephosphorylated by PKA and dephosphorylated at CAD MAPK site late in S phase (Sigoillot et al., 2003). Furthermore, CPS II activity, which catalyzes the initial and rate-limiting step in de novo pyrimidine biosynthesis, is controlled by mitogen-activated protein kinase (MAPK)- and protein kinase A (PKA)-mediated phosphorylation. Changes in CAD enzymatic activity by phosphorylation and dephosphorylation was also observed in rapidly growing cells compared with quiescent cells. Baby hamster kidney cells entering the exponential growth phase exhibited an eightfold increase in pyrimidine biosynthesis, with a fourfold increase in CAD threonine phosphorylation and a 40-fold increase in MAPK activity. In contrast, confluent cells exhibited a twofold increase in CAD phosphoserine modification, a measure of PKA phosphorylation (Sigoillot, Evans, & Guy, 2002). These observations suggest that CAD is activated by MAPK phosphorylation during periods of rapid growth and downregulated by PKA phosphorylation during quiescence (Sigoillot et al., 2002). In normal MCF10A breast cells, pyrimidine biosynthetic pathway was upregulated in the exponential growth phase by MAP kinase phosphorylation of CAD Thr456, and downregulated by P~Thr456 dephosphorylation and PKA phosphorylation of CAD (Sigoillot, Sigoillot, & Guy, 2004).

Enzymes in pyrimidine biosynthetic pathway are also under metabolic control by allosteric effectors (reviewed in (Evans & Guy, 2004)). CAD is regulated by feedback inhibition by the end product uridine 5´-triphosphate (UTP) and allosterically activated by phophoribosyl pyrophosphate (PRPP) (Evans & Guy, 2004). When phosphorylated by MAP kinase or activated by epidermal growth factor, CAD became resistant to feedback inhibition and more sensitive to activation by PRPP (Graves et al., 2000). Increased sensitivity to PRPP and reduced inhibition by UTP in CAD were observed as baby hamster kidney cells approached S phase of the cell cycle. These changes were reversed when cells emerged from S phase (Sigoillot et al., 2003). In vitro and in vivo studies suggest that allosteric regulation of CAD controls the rate of pyrimidine biosynthesis (Evans & Guy, 2004; Sigoillot et al., 2003). Although these activities lie upstream of folate-dependent de novo thymidylate synthesis, their cell cycle regulation provides the needed levels of nucleotide precursor substrate in the form of dUMP to support folate-dependent thymidylate biosynthesis and DNA synthesis during S phase.

2.2.2 |. Dihydrofolate reductase (DHFR)

DHFR expression

DHFR catalyzes the reduction of dihydrofolate (DHF) to tetrahydrofolate (THF), an essential reaction in the de novo thymidylate biosynthesis pathway. DHFR activity is also critical for the reduction of dietary folic acid, a form of folate found in fortified foods and in nutritional supplements, to DHF. DHFR transcript levels are cell cycle regulated. In synchronized human osteosarcoma U2OS cells, DHFR mRNA levels were elevated at the G1/S boundary (Grant et al., 2013). The DHFR gene was expressed at the beginning of S phase in Hela cells and was grouped in the “late DNA replication” cluster by hierarchical clustering analysis (Whitfield et al., 2002). DHFR gene expression was enriched in G2/M phase in synchronized primary human foreskin fibroblasts (Bar-Joseph et al., 2008).

DHFR transcript levels are cell cycle regulated, but there is conflicting evidence regarding the underlying mechanisms. Multiple studies suggest that increased levels of DHFR mRNA during S phase or after serum stimulation was controlled primarily at the level of transcription (Santiago, Collins, & Johnson, 1984; Schilling & Farnham, 1994; Slansky & Farnham, 1996; Wu & Johnson, 1982). In methotrexate-resistant mouse sarcoma cells, Dhfr mRNA levels increased following growth stimulation without a change in Dhfr mRNA half-life (Leys, Crouse, & Kellems, 1984). In methotrexate-resistant mouse 3T6 fibroblasts synchronized by mitotic selection, the transcription rate was low in G1 phase, increased sevenfold at the beginning of S phase, decreased almost immediately thereafter, and remained low throughout the remainder of S and into G2 (Farnham & Schimke, 1985).

The cell cycle regulation of DHFR transcription was achieved by increasing the rate of transcription from a single promoter region (Farnham & Schimke, 1985), mediated through transcription factors and chromatin remodeling (Abali et al., 2008). The hamster Dhfr gene promoter contains consensus binding sites for two eukaryotic transcription factors: Sp1 and E2F. There are four Sp1 cis elements in the mouse Dhfr promoter that play a role in Dhfr transcription (Slansky & Farnham, 1996) in addition to an E2F1 element that is conserved across the human, mouse and hamster Dhfr promoters (Abali et al., 2008). Sp1 binds to the human DHFR promoter throughout the cell cycle in early passage and senescent cells, whereas the E2F binding activity was serum-inducible and was diminished in senescent cells (Good et al., 1996). The E2F family of transcription factors plays an important role in the regulation of gene expression at the G1/S phase transition of the cell cycle. E2F1 expression stimulated the DHFR promoter 22-fold in serum-starved NIH 3T3 cells (Slansky, Li, Kaelin, & Farnham, 1993). However, expression of E2F1 in quiescent cells only minimally induced Dhfr transcription in REF52 cells (DeGregori et al., 1995). The retinoblastoma (RB) tumor suppressor and its family members, p107 and p130, repress E2F activity. There is evidence that E2F-p130 and Sp1-pRB complexes cooperate in repression of the Chinese hamster ovary (CHO) Dhfr gene when cells withdraw from the cell cycle and enter G0 (Chang, Illenye, & Heintz, 2001). In U2OS cells, HDAC1 acts through Sp1 to repress DHFR promoter activity, and the E2F element modulates the activity of SP1 at the DHFR promoter through a cis-acting mechanism (Park et al., 2003). Dhfr expression was induced early in p107−/−; p130−/− mouse embryonic fibroblasts (MEFs) after serum stimulation (Hurford et al., 1997). The mechanism of Sp1 and E2F in regulation of DHFR gene expression has been extensively reviewed elsewhere (Abali et al., 2008).

However, other studies failed to observe cell cycle-dependent regulation of DHFR transcription. One study found Dhfr transcription rates to be invariant throughout the cell cycle, while Dhfr mRNA accumulated as mouse thymocytes progressed though the cell cycle (Feder et al., 1990). In methotrexate-resistant mouse sarcoma cells, Dhfr mRNA levels increase following growth stimulation without a change in Dhfr gene transcription rate. In proliferating cells, most Dhfr transcripts were shown to be converted to mRNA, whereas in quiescent cells, the majority of Dhfr transcripts were rapidly degraded in the nucleus (Leys et al., 1984). Conflicting findings related to DHFR transcriptional regulation through cell cycle may be due to limitations of the nuclear run-on technique and need further investigation (Schilling & Farnham, 1994). Regardless of the mechanism, there is strong evidence that DHFR activity is elevated during S phase to support de novo thymidylate biosynthesis.

DHFR activity exhibits similar cell cycle-dependent patterns as its mRNA levels. DHFR activity was observed to be at its lowest level in G0 and G1 in human lymphoblasts (Pelka-Fleischer, Ruppelt, Wilmanns, Sauer, & Schalhorn, 1987), but increased significantly when quiescence temperature-sensitive cells were stimulated by serum at the permissive temperature (Liu et al., 1985). DHFR steady-state mRNA level, protein synthesis and activity increased twofold as cells progressed from G1 to G2/M in Chinese hamster ovary cells (Feder, Assaraf, Seamer, & Schimke, 1989). DHFR protein has been shown to autoregulate its own translation by binding to DHFR mRNA and inhibiting its translation (Ercikan-Abali et al., 1997). Occupation of the DHFR folate cofactor binding site by methotrexate prevented the interaction of the DHFR protein with its cognate mRNA, thereby relieving translation autoregulation and resulting in increased DHFR levels in cells (Ercikan-Abali et al., 1997). This feedback regulation ensures DHFR enzyme levels do not increase in the absence of available folate cofactor availability.

DHFR subcellular localization

HeLa cells expressing a GFP-DHFR fusion protein provided evidence for cell cycle-dependent subcellular localization of DHFR. The GFP-DHFR fusion protein localized to the nucleus during S and G2/M phases but not in G1 phase (Anderson, Woeller, et al., 2012). Nuclear localization may be small ubiquitin-like modifier (SUMO) dependent. DHFR was shown to be modified by SUMO-1 in vitro (Anderson, Woeller, & Stover, 2007) and was also identified as a target for SUMO2 modification in proteome-wide mass spectrometry analysis (Tammsalu et al., 2014). Potential SUMOylation modification sites were identified through mass spectrometry analysis and through sequence alignments from various species, but these observations require experimental verification (Anderson et al., 2007; Hendriks, D’Souza, Chang, Mann, & Vertegaal, 2015; Tammsalu et al., 2014).

2.2.3 |. Serine hydroxymethyltransferase (SHMT1 and SHMT2α)

SHMT1 expression

There are two cytosolic/nuclear isozymes of SHMT encoded by distinct genes; SHMT1 and SHMT2α and these two enzymes are functionally redundant (Anderson & Stover, 2009). They catalyze the reversible conversion of serine and THF to glycine and 5,10-methylene-THF. One study found that SHMT1 mRNA levels were elevated in G1/S phase of the cell cycle in synchronized U2OS cells (Grant et al., 2013), whereas in another study, SHMT1 protein levels were elevated during S phase in HeLa cells blocked with hydroxyurea without changes in SHMT1 mRNA level (Anderson, Eom, & Stover, 2012; Field et al., 2014). A significant increase in SHMT activity was observed in lymphocytes when stimulated to proliferate by phytohemagglutinin treatment (Thorndike, Pelliniemi, & Beck, 1979).

SHMT subcellular localization

SHMT1 and SHMT2α and their cytosolic/nuclear localization are cell cycle regulated. The SHMT2 gene encodes an SHMT2 protein that localizes exclusively to the mitochondria, whereas it also expresses a SHMT2α isozyme that localizes to the cytosol and translocates to the nucleus during S phase (Anderson & Stover, 2009; Anderson, Woeller, et al., 2012). SHMT1 and SHMT2α are modified by SUMO during S phase of the cell cycle, and SUMOylated SHMT1 is enriched and exhibits primarily nuclear localization in HeLa cells (Woeller, Anderson, Szebenyi, & Stover, 2007). SHMT1 was localized primarily to the cytoplasm at G1/stationary phase of the cell cycle and to the cytoplasm, nuclear periphery, and nucleus in the S phase and G2/M phase of the cell cycle in blocked MCF-7 cells (Woeller et al., 2007). Evidence suggests that SUMOylation of SHMT1 occurs at the nuclear pore and is linked to its nuclear import, as mutation of either SHMT1 Lys 38 or Lys 39 to arginine impaired in vitro SUMOylation by SUMO1 and nuclear localization of the mutant protein. Ran-binding protein 2, a nuclear periphery/nuclear pore protein which possesses E3 SUMO ligase activity, also plays a role in SHMT1 nuclear localization (Woeller et al., 2007). It is likely that the other enzymes in the de novo dTMP synthesis pathway, DHFR and TYMS, are compartmentalized in the nucleus during S phase through similar mechanisms.

Interestingly, SHMT1 is also ubiquitinated at Lys39 and is subject to ubiquitin-dependent degradation. Ub-SHMT1 was at the lowest level at S phase of the cell cycle. Lys-63 polyubiquitination of SHMT1 was enriched in the nuclear fraction in S and G2/M phase, but absent in G1 in both nuclear and cytosolic fractions. Mutation of the ubiquitination site increases SHMT1 stability (Anderson, Eom, & Stover, 2012). In HeLa cells, Ubc13-mediated ubiquitination of SHMT1 is required for nuclear export and nuclear stability, whereas Ubc9-mediated modification with Sumo2/3 leads to SHMT1 degradation within the nucleus (Anderson, Eom, & Stover, 2012).

2.2.4 |. Thymidylate synthase (TYMS)

TYMS expression

TYMS catalyzes the reductive methylation of deoxyuridylate (dUMP) to deoxythymidylate (dTMP) using 5,10-methylene-THF as a cofactor. DTMP is further phosphorylated to dTTP for DNA synthesis. TYMS activity and protein levels are subject to cell cycle variation. Very low-TYMS enzyme activity in G0 and G1 phase has been observed in human lymphoblasts with significant higher levels in S and G2/M phases (Pelka-Fleischer et al., 1987; Pelka-Fleischer, Fleischer, Wilmanns, Sauer, & Schalhorn, 1989). Elevations in TYMS protein levels and activity are associated with the onset of S phase when cells were growth arrested and then allowed to reenter the proliferating state (reviewed in (Dolnick, 2003)). In human diploid fibroblasts synchronized by serum starvation, TYMS protein and mRNA level increased through post-transcriptional regulation (Ayusawa et al., 1986). In MCF-7 cells synchronized with lovastatin, TYMS protein level is tightly regulated in the cell cycle with the peak of TYMS level coinciding with that of S phase, with no change in the level of TYMS mRNA level (Keyomarsi, Samet, Molnar, & Pardee, 1993). Similarly, other studies have shown that TYMS cell cycle regulation occurs through intron splicing signals, with very little change in rate of transcription during G1-S phase transition in serum stimulated mouse 3T6 fibroblasts (Ash, Liao, Ke, & Johnson, 1995; Ke, Ash, & Johnson, 1996).

Other evidence suggests that the TYMS gene is regulated transcriptionally and post-transcriptionally (reviewed in (Chu & Allegra, 1996; Dolnick, 2003)). HCT116 cells synchronized by serum starvation exhibit increased TYMS mRNA levels during G1, followed by an increase in TYMS protein levels which did not decrease following S phase completion (Le François, Maroun, & Birnboim, 2007). Tyms mRNA levels increased 20- to 40-fold as mouse fibroblasts progressed from a resting to late S phase, as a result of increased transcription rates and other post-transcriptional regulation (Jenh, Geyer, & Johnson, 1985). In synchronized HeLa cells, analysis of transcript levels and hierarchical clustering of the expression patterns classified TYMS in the “late DNA replication” cluster, together with other genes in nucleotide metabolism, including DHFR, ribonucleotide reductase (RRM1 and RRM2), and additional DNA replication, repair and recombination genes (Whitfield et al., 2002). Interestingly, TYMS expression was shown to be downregulated during metastatic HO-1 human melanoma cells differentiation (Huynh et al., 2009). In cells that have not been subjected to cell cycle arrest followed by release, analyses of TYMS protein and mRNA levels lead to conflicting results. Several studies found no variation in TYMS protein or TYMS mRNA with S phase (reviewed in (Dolnick, 2003)). On the contrary, TYMS mRNA has been observed to accumulate as thymocytes progress through the cell cycle (Feder et al., 1990).

The TYMS gene is regulated by several transcription factors. Transcription factor Late SV40 Factor (LSF) induces TYMS gene at G1/S transition in growth-stimulated cells through binding to sites within the TYMS promoter and intronic regions (Powell et al., 2000). The TYMS gene is also targeted by the transcription factor Forkhead Box M1 (FOXM1) (Huynh et al., 2011). Chromatin immunoprecipitation followed by high-throughput sequencing (ChIP-seq) in U2OS cells identified TYMS gene as a target for FOXM1, which strongly activates promoters of G2/M phase genes and weakly activates those induced in S phase (Grant et al., 2013). Tyms gene expression has also been shown to be regulated by E2F in MEFs (Ishida et al., 2001). Tyms expression was derepressed in G0 and G1 in MEFs lacking both p107 and p130, two E2F activity controlling proteins (Hurford et al., 1997). DNA microarray analyses demonstrated that Tyms gene expression is upregulated by E2F-1 or E2F-3, along with the functionally-related enzymes Uracil-DNA glycosylase (UNG), proliferating cell nuclear antigen, and dUTP nucleotidohydrolase (Polager et al., 2002). Tyms gene transcription was induced by E2F1 in quiescent REF52 cells infected with E2F1 cDNA containing recombinant adenovirus (DeGregori et al., 1995). E2F motifs in rat Tyms promoter region were found to be necessary for increased Tyms expression during the G1 to S transition in serum stimulated cells (Lee & Johnson, 2000). However, adenovirus-mediated over-expression of E2F1 and cyclin E in three human cell lines had no effect on TYMS expression (Le François et al., 2007). As seen for DHFR, TYMS protein autoregulates its synthesis by binding to its own mRNA with high affinity which represses translation (reviewed in (Chu & Allegra, 1996)).

TYMS subcellular localization

HeLa cells expressing a GFP-TYMS fusion protein showed nuclear TYMS localization during S and G2/M phases but not in G1 (Anderson, Woeller, et al., 2012). TYMS is modified by SUMO1 in vitro (Anderson et al., 2007), and was also identified as a target for SUMO2 modification in system-wide mass spectrometry analysis (Hendriks et al., 2015; Tammsalu et al., 2014). Several putative SUMOylation modification sites were identified through mass spectrometry analysis (Anderson et al., 2007; Hendriks et al., 2015; Tammsalu et al., 2014). Collectively, there is strong evidence for cell cycle regulation of the enzymes that constitute the de novo dTMP synthesis pathway, with elevated protein levels and nuclear localization of the pathway during S phase.

2.3 |. dTMP salvage pathway

2.3.1 |. TK expression

Thymidine kinase 1 (TK1) catalyzes the conversion of thymidine to dTMP in pyrimidine salvage pathway synthesis, and its enzyme activity varies throughout the cell cycle and reflects the levels of TK1 protein (Sherley & Kelly, 1988). TK1 activity was shown to vary through the cell cycle in synchronized mouse fibroblasts and synchronized human KB cells (Bello, 1974; Littlefield, 1966). Very little or no activity of TK1 was observed in G0 and G1 in human lymphoblasts (Pelka-Fleischer et al., 1987; Pelka-Fleischer et al., 1989; Piper, Tattersall, & Fox, 1980), with a higher level of TK1 activity observed in S and G2/M (Pelka-Fleischer et al., 1989), with the maximum activity observed after the peak of DNA synthesis (Piper et al., 1980). In contrast to increased TK1 activity in S and G2/M phases, enzymes involved in thymidine and thymidylate catabolism, including dihydrothymine dehydrogenase, thymidine phosphorylase and dTMP phosphatase, have constant levels of activity throughout the cell cycle in synchronized Human B lymphocytes (Piper et al., 1980).

Both TK1 mRNA and protein levels were shown to be upregulated in rapidly dividing cells. Tk1 mRNA levels increased more than 20-fold in S phase compared to G0-phase of the cell cycle in BALB/c 3T3 cells (Coppock & Pardee, 1987). Proliferating cells exhibited higher levels of TK1 mRNA and protein compared to slowly growing, quiescent, or terminally differentiated cells. The levels of TK1 mRNA and enzyme activity increased significantly when quiescent cells were stimulated by serum or adenovirus infection (Lewis & Matkovich, 1986; Liu, Baserga, & Mercer, 1985; Liu, Gibson, et al., 1985).

It is not clear how TK1 protein levels are regulated throughout the cell cycle. TK1 mRNA levels were increased in senescent human diploid fibroblasts WI38 when serum stimulated, without increased incorporation of 3H thymidine (Kaczmarek, 1986) into DNA, indicating that TK1 activity was not increased with elevated transcript levels. However, increased TK1 activity depended on the availability of TK1 mRNA in BALB/c 3T3 cells (Coppock & Pardee, 1987). Pulse labeling experiments showed that TK1 protein synthesis was elevated 10-fold in S phase compared to G1 phase, suggesting that the efficiency of translation of TK1 mRNA increased (Sherley & Kelly, 1988). TK1 protein synthesis ends with a post-transcriptional block at about the time human KB cells begin mitosis (Bello, 1974). Conflicting data exist with regard to TK1 protein stability throughout the cell cycle. TK1 protein stability changes during the cell cycle in some circumstances, as the stability of TK1 protein was shown to decrease upon cell division in HeLa cells (Sherley & Kelly, 1988). However, in human KB cells, the stability of the enzyme did not change significantly throughout the cell cycle, suggesting that enzyme activity is determined by rate of enzyme synthesis rather than enzyme degradation (Bello, 1974). Interestingly, deletion of the carboxyl-terminal 40 amino acids or fusion of beta-galactosidase to the carboxylterminus of human TK1 completely abolished cell cycle regulation and stabilized the protein throughout the cell cycle (Kauffman & Kelly, 1991).

TK1 mRNA levels are controlled by both rates of gene transcription and mRNA degradation, and subject to cell cycle regulation. In growth stimulated CV1 African Green Monkey kidney cells, a large increase in TK1 mRNA levels and a relatively small increase in transcription rates were observed, suggesting that TK1 gene expression is controlled at both a transcriptional and post-transcriptional level (Stewart, Ito, & Conrad, 1987). In BALB/c 3T3 cells, the rate of run-on Tk1 transcription increased twofold to fourfold during S phase compared to G0 phase (Coppock & Pardee, 1987). The half-life of Tk1 mRNA was longer in S and M phases then in quiescence (Coppock & Pardee, 1987).

The mechanism of TK1 transcriptional cell cycle regulation remains elusive. In CV1 African Green Monkey kidney cells, the TK1 cDNA alone is sufficient to encode cell cycle regulated expression (Stewart et al., 1987). However, Travali et al. showed that the Tk1 promoter has also an important role in cell cycle regulation of TK1 mRNA levels (Travali, Lipson, Jaskulski, Lauret, & Baserga, 1988). Tk1 gene transcription was regulated by E2F in MEFs (Ishida et al., 2001), however, Tk1 expression was not affected in MEFs lacking E2F regulating proteins Rb, p107, p130, or both 107, and p130 (Hurford et al., 1997). Similar to the TYMS gene, the TK1 gene is also bound by transcription factor FOXM1 in U2OS and HeLa cells (Grant et al., 2013).

2.3.2 |. TK subcellular localization

Like the enzymes that comprise the de novo dTMP synthesis pathway, TK1 translocates into the nucleus during S phase of the cell cycle, providing dTMP at site of DNA replication. In Chinese hamster cells during S phase, TK1 co-fractionated with DNA polymerase, NDP kinase, RNR, and the de novo dTMP synthesis pathway enzymes, including DHFR and TYMS (Noguchi, Prem veer Reddy, & Pardee, 1983). The mechanism for TK1 nuclear localization during S phase remains unknown.

2.4 |. Purine synthesis

Human cells produce purines in the cytosol through both salvage and de novo biosynthesis pathways. The purine nucleotide salvage pathway includes a one-step conversion of hypoxanthine to inosine monophosphate (IMP) (Yamaoka et al., 1997). IMP is further converted into adenosine and guanosine nucleotides. The more energy consuming de novo pathway consists of 10 chemical reactions that convert phosphoribosyl pyrophosphate (PRPP) into IMP, utilizing the 1C from two 10-formylTHF during the process (the formyl group of 10-formylTHF is incorporated as the #2 and #8 carbons of the purine ring). In order to conserve energy, purine nucleotides are synthesized preferentially through salvage pathway when hypoxanthine is available in most cells (Kondo et al., 2000). However, de novo purine synthesis, rather than purine salvage synthesis or pyrimidine synthesis, limits CHO cell growth rate (Kondo et al., 2000).

2.4.1 |. Expression of enzymes in purine synthesis pathway

Purine nucleotide synthesis rates are growth dependent in rat livers (Jackson et al., 1980). De novo purine biosynthesis was shown to be upregulated in serum stimulated 3T6 fibroblasts, with increased levels of PRPP and formylglycinamide ribonucleotide (biosynthetic intermediates in the de novo purine synthesis pathway) (Smith & Buchanan, 1979). De novo and salvage purine synthesis pathway increased 5- and 3.3-fold, respectively, as cells progressed from mid-G1 phase to early S phase (Fridman et al., 2013). In CHO K1 cells synchronized by serum deprivation, amidophosphoribosyltransferase (ATase) and hypoxanthine-guanine phosphoribosyltransferase (HPRT), which are key regulatory enzymes of the de novo and salvage pathways of purine synthesis respectively, showed distinct expression changes throughout the cell cycle. ATase activity increased from late G1 phase to the S phase, whereas HPRT activity was nearly constant during throughout the cell cycle (Kondo et al., 2000). ATase and aminoimidazole ribonucleotide carboxylase (an enzyme in de novo purine synthesis pathway) mRNA levels increased approximately fivefold to sixfold in G1/S phase of the cell cycle compare to G0 in synchronized rat 3Y1 fibroblasts (Iwahana et al., 1995). MTORC1 activity, which is important for S and G2 progression (Robitaille et al., 2013) and M phase entry (review in (Proud, 2010)), increased metabolic flux through the de novo purine synthesis pathway and adjusted purine nucleotide pools available for nucleic acid synthesis in various mouse and human cells (Ben-Sahra, Hoxhaj, Ricoult, Asara, & Manning, 2016). Similarly, purine synthesis was stimulated by mTORC1 through activating transcription factor 4 (ATF4) in both normal and cancer cells (Ben-Sahra et al., 2016).

2.4.2 |. Subcellular localization of enzymes in the do novo purine synthesis pathway

The six proteins that comprise the de novo purine synthesis pathway were shown to cluster in the cytoplasm and form a multienzyme complex known as the “purinosome” in HeLa cells under purine depletion (An, Kumar, Sheets, & Benkovic, 2008). Purinosome formation exhibited cell cycle-dependence in HeLa cells cultured in purine-depleted conditions, with the highest number of cells with purinosomes observed during G1 phase (Chan et al., 2015). A similar observation was made in Lesch–Nyhan disease (LND) fibroblast cells that are deficient in hypoxanthine-guanine phosphoribosyl transferase (HGPRT), an enzyme in the purine salvage pathway. However, elevated levels of purinosomes in S and G2/M phases were also observed, probably due to the fact that LND cells rely primarily on the de novo pathway to synthesize purine nucleotides (Chan et al., 2015).

2.5 |. Methenyltetrahydrofolate synthetase (MTHFS)

2.5.1 |. MTHFS expression

MTHFS catalyzes the irreversible and ATP-dependent conversion of 5-formylTHF (a storage form of THF cofactors) to 5,10-methenylTHF. 5-formylTHF is an inhibitor of SHMT (Girgis, Suh, Jolivet, & Stover, 1997) and AICAR transformylase (Bertrand & Jolivet, 1989) (an enzyme in the de novo purine synthesis pathway). MTHFS protein activity is inhibited by 10-formylTHF, which exists in chemical equilibrium with 5,10-methenylTHF (Field et al., 2006).

MTHFS gene expression was shown to be significantly higher in G1/S phase of the cell cycle in synchronized HacaT human keratinocytes (Peña-Diaz et al., 2013). Although the MTHFS gene has been observed to bind the transcription factor FOXM1 (Grant et al., 2013), the mechanism of cell cycle regulated gene expression of MTHFS remains unknown.

2.5.2 |. MTHFS subcellular localization

MTHFS cellular localization has been shown to be cell cycle dependent. A GFP-MTHFS fusion protein colocalized with purinosomes in purine-depleted media in Hela cells, suggesting MTHFS is a component of the purinosome (Field, Anderson, & Stover, 2011). It has been proposed that MTHFS delivers 10-formylTHF cofactors to the purinosome, as MTHFS levels influence rates of de novo purine biosynthesis (Field et al., 2011). SUMOylation was shown to be necessary for localization of GFP-MTHFS to the purinosome. Recombinant MTHFS is modified by SUMO-1 both in vitro and in Hela cells, and mutation of the MTHFS SUMO1 modification sites K140 and K190 ablated the localization of a GFP-MTHFS fusion protein to the purinosome in HeLa cells cultured in purine-deficient conditions (Field et al., 2011).

2.6 |. Homocysteine remethylation pathway

2.6.1 |. Methylenetetrahydrofolate reductase (MTHFR)

There is limited evidence for cell cycle regulation of MTHFR, an enzyme that catalyzes the conversion of 5,10-methylene-THF to 5-methylTHF (a required cofactor for homocysteine remethylation to methionine). MTHFR is phosphorylated at T34 by CDK1/cyclin B1 (Zhu et al., 2014) which inhibits its catalytic activity and this phosphorylation peaked during mitosis in Hela cells (Zhu et al., 2014).

2.6.2 |. Methionine synthase (MTR)

MTR is a folate- and vitamin B12-dependent enzyme that catalyzes the conversion of homocysteine into methionine, utilizing a methyl group from 5-methylTHF which is transferred to homocysteine through a methyl-cobalamin intermediate. Resting human peripheral blood lymphocytes do not efficiently take up vitamin B12 in the form of cobalamin (CbI); increased CbI uptake correlates with active cell division and DNA synthesis through increased receptor activity (Hall, 1984). The MTR mRNA level is elevated in S phase of the cell cycle in synchronized U2OS cells (Grant et al., 2013). The lowest level of MTR activity was observed during G0 and G1 in human lymphoblasts (Pelka-Fleischer et al., 1987). Another study found MTR activity correlated with cell division and DNA synthesis in human lymphocytes (Hall, 1984).

2.7 |. Other FOCM related enzymes and transporters

2.7.1 |. Folate transporters

Folates are transported into cells through membrane-bound folate receptor alpha (FR-α), the reduced folate carrier (RFC), and the proton-coupled folate transporter (PCFT). Folate uptake is closely correlated with cell growth and S phase of the cell cycle. FR-α expression has been shown to be cell cycle-dependent with the highest expression levels in S phase (Tang et al., 2015). FR-α function has been observed to decrease as cellular growth slowed in JAR, Caco-2 and MA-104 cell lines (Doucette & Stevens, 2001). In primary human trophoblast cells, inhibition of mTORC1 or mTORC2 markedly decreased basal folate uptake and decreased the plasma membrane expression of FR-α and RFC transporter isoforms without changes in global protein expression levels (Rosario, Powell, & Jansson, 2016).

2.7.2 |. Ribonucleotide reductase

Ribonucleotide reductase (RNR) converts ribonucleotides to deoxyribonucleotides, an activity that is essential for de novo dTMP synthesis by generating the precursor substrate dUMP. RNR consists of two subunits, RRM1 and RRM2. RRM1 protein levels remained constant throughout the cell cycle in bovine kidney MDBK cells (Engström et al., 1985), whereas RRM2 protein levels increased as cells passed from G1 to S (Eriksson, Gräslund, Skog, Thelander, & Tribukait, 1984). During G2, RRM2 was degraded, which maintained balanced dTNP pools and genome stability (D’Angiolella et al., 2012). RRM2 interacts with cycline F and its protein level is tightly controlled by two different ubiquitin ligases in the G1 and G2 phases of the cell cycle (D’Angiolella, Esencay, & Pagano, 2013). RNR activity is highest during S phase in eukaryotes, when the requirement for dNTPs is the highest (reviewed in (Nordlund & Reichard, 2006)).

RNR gene expression has been shown to be cell cycle regulated (Ishida et al., 2001). Both Rrm1 and Rrm2 transcripts were very low or undetectable in G0 and G1, but increased as cells progressed into S phase, and then declined in G2/M phase in hydroxyurea-resistant mouse mammary tumor TA3 cells (Björklund, Skog, Tribukait, & Thelander, 1990). The regulation of RNR transcription is not well understood. Rrm1 and Rrm2 transcription rates are invariant throughout the cell cycle in mouse thymocytes, whereas Rrm2 mRNA levels but not Rrm1 exhibited cyclic regulation in mouse thymocytes (Feder et al., 1990). Another study indicated that Rrm1 gene expression is mainly regulated at the transcriptional level during the cell cycle (Johansson, Hjortsberg, & Thelander, 1998). Rrm1 and Rrm2 transcription was induced by E2F1 expression in quiescent cells (DeGregori et al., 1995) and in synchronized MEFs (Ishida et al., 2001). Rrm2 expression was derepressed in G0 and G1 in MEFs lacking both p107 and p130, which control E2F activity (Hurford et al., 1997). S phase-specific transcription of the mouse Rrm2 gene is dependent on one proximal promoter repressive element that binds E2F4, and mutation of the E2F binding site leads to premature promoter activation in G1 (Chabes et al., 2004).

3 |. FOCM IN THE NUCLEUS

3.1 |. Nuclear co-localization of enzymes in the de novo dTMP synthesis pathway

During S and G2/M phases of the cell cycle or following DNA damage, the enzymes of the dTMP synthesis pathway, including SHMT1, SHMT2α, TYMS, DHFR, and MTHFD1 are SUMOylated and translocate into the nucleus for DNA replication or repair (Anderson, Woeller, et al., 2012). This pathway forms a multienzyme complex that is associated with the nuclear lamina and other enzymes of the DNA replication machinery (Anderson et al., 2007; Anderson & Stover, 2009; Anderson, Woeller, et al., 2012; Woeller et al., 2007). SHMT1 and SHMT2α independently serve as scaffold proteins that are essential for dTMP synthesis complex formation at sites of DNA replication (Anderson, Woeller, et al., 2012). Interestingly, in BHK cells transfected with a fluorescent CAD fusion protein, a significant fraction of CAD translocates in the nucleus when cells enter S phase of the cell cycle (Sigoillot et al., 2003). The role of CAD nuclear localization remains unknown (Evans & Guy, 2004). Similarly, RNR has been observed to localize at DNA damage sites to allow production of dNTPs at sites of DNA repair (Niida et al., 2010), indicating that the nucleus may synthesize thymidylate de novo completely within the nuclear compartment in some cells.

The co-localization of the enzymes in the de novo dTMP synthesis pathway suggests that the spatial organization of the pathway enzymes play an important role in meeting cellular dTMP demand during DNA replication in S phase. The formation of the dTMP synthesis complex at sites of DNA replication potentially enables much more efficient catalysis through substrate channeling, which offers kinetic advantages over free diffusion of folate cofactors within the bulk solvent (Misselbeck et al., 2017).

3.2 |. Uracil-DNA glycosylases (UNG)

Folate deficiency and impaired dTMP synthesis leads to uracil misincorporation into DNA and chromosome instability and breakage. UNG excises uracil residues from DNA. Specifically, UNG cleaves the N-glycosylic bond and initiates the base-excision repair pathway. UNG gene expression is enriched in the G1/S and G2 phases of the cell cycle in synchronized primary human foreskin fibroblasts (Bar-Joseph et al., 2008).

4 |. FOCM IN THE MITOCHONDRIA

1C units are generated in mitochondria through catabolism of serine, glycine, histidine, betaine, dimethylglycine and sarcosine, and are released into the cytosol in the form of formate (Tibbetts & Appling, 2010). Mammalian mitochondria also contain a de novo dTMP synthesis pathway, with SHMT2 playing an essential role in limiting uracil misincorporation into mitochondrial DNA (Anderson, Quintero, & Stover, 2011). Confocal microscopy demonstrated that fluorescent fusion proteins of DHFR2 and TYMS localized to the mitochondria in Hela cells. Endogenous DHFR2, SHMT2 and TYMS have also been shown to localize to the mitochondrial matrix and the inner membrane of the mitochondria in fractionated HepG2 mitochondria (Anderson et al., 2011).

4.1 |. Methylenetetrahydrofolate dehydrogenase 2

Methylenetetrahydrofolate dehydrogenase (MTHFD2) is a nuclear-encoded mitochondrial bifunctional enzyme that has methylene-THF dehydrogenase and methenylTHF cyclohydrolase activities. The multifunctional enzyme reversibly converts 5,10-methylene-THF, 5,10-methenylTHF and 10-formyl THF. The formyl group of 10-formyl THF is liberated as formate, thereby generating THF via the enzyme MTHFD1L. MTHFD2 is also found in the nucleus, though its function within the nucleus has not been elucidated (Gustafsson Sheppard et al., 2015). MTHFD2 has recently been shown to co-localize with newly synthesized DNA in the nuclei of U251, HeLa, and HCT116 cells (Gustafsson Sheppard et al., 2015). MTHFD2 gene expression is upregulated in S phase of the cell cycle in U2OS cells (Grant et al., 2013). MTHFD2 expression was stimulated by mTORC1 through activating transcription factor 4 in both normal and cancer cells (Ben-Sahra et al., 2016). MTHFD2 over-expression also increased HCT116 proliferation rates, even when MTHFD2 lacking functional dehydrogenase activity was expressed (Gustafsson Sheppard et al., 2015). This observation, taken together with the assumption that MTHFD2 dehydrogenase activity would be expected to decrease synthesis of 5,10-methylene-THF (or oppose the activity of MTHFD1 dehydrogenase activity), suggests that the function of MTHFD2 within the nucleus is distinct from its catalytic activity.

4.2 |. Serine hydroxymethyltransferase 2

The SHMT2 gene encodes two transcripts, a mitochondrial SHMT2 isozyme and a cytoplasmic/nuclear SHMT2α isozyme. SHMT2 expression is significantly enriched in G2/M phase of the cell cycle in synchronized HacaT human keratinocytes (Peña-Diaz et al., 2013), whereas SHMT2 mRNA levels in MCF-7 cells did not vary during the cell cycle (Stover et al., 1997). SHMT2 expression is predicted to be regulated by transcription factor E2F and Sp1 (Lane & Fan, 2015).

4.3 |. Compartmentalization of folate derivatives

Folate derivatives are compartmentalized within the cell. Mitochondria contain roughly 40% of total cellular folate (Lin, Huang, & Shane, 1993; Shin, Chan, Vidal, Brody, & Stokstad, 1976), while the nuclear compartment contains about 10% cellular folate (Shin et al., 1976), leaving about 50% of cellular folate within the cytosol. Neither total cellular folate levels nor nuclear folate levels varied as a function of cell cycle in MCF-7 cells (Field et al., 2014). Interestingly, nuclear folate levels resisted depletion during S phase in MCF-7 cells cultured in folate-depleted media (Field et al., 2014). In mice consuming folic acid-deficient diets for 6 weeks, there was a 50% decrease in liver whole-cell folate concentration compared to animals consuming control diets containing folic acid, but nuclear folate levels resisted depletion (Field et al., 2014). These data suggest that nuclear folate levels are maintained to protect de novo dTMP synthesis during S phase and in response to folate deficiency. However, protecting nuclear folate pools and dTMP synthesis occurs at the expense of cytosolic folate pools and homocysteine remethylation (Field et al., 2015).

5 |. CONCLUSIONS

Nutrition and genetic epidemiological studies and/or randomized controlled trials implicate impaired folate status and FOCM in several pathologies, including megaloblastic anemia, neural tube defects, neurodegenerative disease, and various types of cancer (Crott, Liu, Choi, & Mason, 2007; Friso et al., 2002; Friso, Choi, Dolnikowski, & Selhub, 2002; Huang, Sloan, & Boerkoel, 2003; Jaenisch & Bird, 2003; Suh, Herbig, & Stover, 2001; Tufi et al., 2014). However, the causal pathways and mechanisms underlying these pathologies remain unclear due to the interconnectedness of these pathways (Misselbeck et al., 2017). Key to understanding the etiology of folate-associated pathologies will be elucidating the regulation of the pathways within FOCM, and how decisions are made to partition one-carbon units and other intermediates among the pathways, as well as the partitioning of folate cofactors which are a limiting resource for all pathways (Suh et al., 2001). Cell cycle regulation of FOCM, which is necessary considering its role in DNA synthesis, provides a mechanism to ensure cellular needs are met by assigning priority to individual pathways within the network based on temporal needs, especially deoxyribonucleotide biosynthesis. Evidence for cell cycle regulation of homocysteine remethylation and mitochondrial one-carbon metabolism is much more limited.

There is clear evidence that folate-dependent and folate independent purine and thymine deoxyribonucleotide synthesis are regulated by cell cycle. This regulation involves both an increase in enzyme activity through increased expression and enzyme activity, as well as formation of multienzyme complexes. However, the strategies for meeting cellular demands differ among the pathways. Thymidylate synthesis occurs in the nucleus at sites of DNA synthesis, with both increased expression of the pathway and nuclear import occurring at the S and G2M phases. Nuclear dTMP synthesis separates this pathway from FOCM in the cytosol, eliminating competition for folate cofactors with de novo purine biosynthesis and homocysteine remethylation, assuming that folate nuclear and cytosolic folate cofactors are not shared (Field et al., 2014).

De novo purine biosynthesis and homocysteine remethylation both occur in the cytoplasm and are poised to compete for cofactor availability. Both of these pathways rely on MTHFD1 and cellular formate to provide folate-activated 1C units. De novo purine biosynthesis exhibits cell cycle regulation at the level of multienzyme complex formation, but less information is available concerning cell cycle regulation of enzyme levels. If purinosome formation, which forms at G1 and extends into S phase, is critical to meet cellular purine nucleotide needs, this may suggest that homocysteine remethylation may be most vulnerable to decreased activity during times of purinosome formation.

Understanding how metabolic decisions are made as a function of cell cycle, both in terms of enzyme expression and localization, will shed light on the regulation of FOCM network and the role of disrupted FOCM in disease pathogenesis.

ACKNOWLEDGMENT

This work was support by funding from PHS R37DK58144 and 3R01 HD059120-08S1.

Funding information

PHS, Grant/Award Number: R37DK58144 and 3R01 HD059120-08S1

Footnotes

CONFLICT OF INTEREST

The authors have declared no conflicts of interest for this article.

REFERENCE

  1. Abali EE, Skacel NE, Celikkaya H, & Hsieh Y. (2008). Vitamins and hormones (Vol. 79, pp. 267–292). [DOI] [PubMed] [Google Scholar]
  2. An S, Kumar R, Sheets ED, & Benkovic SJ (2008). Reversible compartmentalization of de novo purine biosynthetic complexes in living cells. Science, 320, 103–106. [DOI] [PubMed] [Google Scholar]
  3. Anderson DD, Eom JY, & Stover PJ (2012). Competition between sumoylation and ubiquitination of serine hydroxymethyltransferase 1 determines its nuclear localization and its accumulation in the nucleus. The Journal of Biological Chemistry, 287, 4790–4799. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Anderson DD, Quintero CM, & Stover PJ (2011). Identification of a de novo thymidylate biosynthesis pathway in mammalian mitochondria. Proceedings of the National Academy of Sciences of the United States of America, 108, 15163–15168. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Anderson DD, & Stover PJ (2009). SHMT1 and SHMT2 are functionally redundant in nuclear De novo thymidylate biosynthesis. PLoS One, 4, e5839. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Anderson DD, Woeller CF, Chiang EP, Shane B, & Stover PJ (2012). Serine hydroxymethyltransferase anchors de Novo thymidylate synthesis pathway to nuclear lamina for DNA synthesis. The Journal of Biological Chemistry, 287, 7051–7062. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Anderson DD, Woeller CF, & Stover PJ (2007). Small ubiquitin-like modifier-1 (SUMO-1) modification of thymidylate synthase and dihydrofolate reductase. Clinical Chemistry and Laboratory Medicine, 45, 1760–1763. [DOI] [PubMed] [Google Scholar]
  8. Ash J, Liao WC, Ke Y, & Johnson LF (1995). Regulation of mouse thymidylate synthase gene expression in growth-stimulated cells: Upstream S phase control elements are indistinguishable from the essential promoter elements. Nucleic Acids Research, 23, 4649–4656. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Ayusawa D, Shimizu K, Koyama H, Kaneda S, Takeishi K, & Seno T. (1986). Cell-cycle-directed regulation of thymidylate synthase messenger RNA in human diploid fibroblasts stimulated to proliferate. Journal of Molecular Biology, 190, 559–567. [DOI] [PubMed] [Google Scholar]
  10. Bar-Joseph Z, Siegfried Z, Brandeis M, Brors B, Lu Y, Eils R, … Simon I. (2008). Genome-wide transcriptional analysis of the human cell cycle identifies genes differentially regulated in normal and cancer cells. Proceedings of the National Academy of Sciences of the United States of America, 105, 955–960. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Bello LJ (1974). Regulation of thymidine kinase synthesis in human cells. Experimental Cell Research, 89, 263–274. [DOI] [PubMed] [Google Scholar]
  12. Ben-Sahra I, Howell JJ, Asara JM, & Manning BD (2013). Stimulation of de novo pyrimidine synthesis by growth signaling through mTOR and S6K1. Science, 339, 1323–1328. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Ben-Sahra I, Hoxhaj G, Ricoult SJH, Asara JM, & Manning BD (2016). mTORC1 induces purine synthesis through control of the mitochondrial tetrahydrofolate cycle. Science, 351, 728–733. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Bertrand R, & Jolivet J. (1989). Methenyltetrahydrofolate synthetase prevents the inhibition of phosphoribosyl 5-aminoimidazole 4-carboxamide ribonucleotide formyltransferase by 5-formyltetrahydrofolate polyglutamates. The Journal of Biological Chemistry, 264, 8843–8846. [PubMed] [Google Scholar]
  15. Björklund S, Skog S, Tribukait B, & Thelander L. (1990). S-phase-specific expression of mammalian ribonucleotide reductase R1 and R2 subunit mRNAs. Biochemistry, 29, 5452–5458. [DOI] [PubMed] [Google Scholar]
  16. Boyd KE, & Farnham PJ (1997). Myc versus USF: Discrimination at the cad gene is determined by core promoter elements. Molecular and Cellular Biology, 17, 2529–2537. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Chabes AL, Björklund S, & Thelander L. (2004). S Phase-specific transcription of the mouse ribonucleotide reductase R2 gene requires both a proximal repressive E2F-binding site and an upstream promoter activating region. The Journal of Biological Chemistry, 279, 10796–10807. [DOI] [PubMed] [Google Scholar]
  18. Chan CY, Zhao H, Pugh RJ, Pedley AM, French J, Jones SA, … Benkovic SJ (2015). Purinosome formation as a function of the cell cycle. Proceedings of the National Academy of Sciences of the United States of America, 112, 1368–1373. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Chang YC, Illenye S, & Heintz NH (2001). Cooperation of E2F-p130 and Sp1-pRb complexes in repression of the Chinese hamster dhfr gene. Molecular and Cellular Biology, 21, 1121–1131. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Chen K-F, Lai Y-Y, Sun HS, & Tsai S-J (2005). Transcriptional repression of human cad gene by hypoxia inducible factor-1alpha. Nucleic Acids Research, 33, 5190–5198. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Chu E, & Allegra CJ (1996). The role of thymidylate synthase as an RNA binding protein. BioEssays, 18, 191–198. [DOI] [PubMed] [Google Scholar]
  22. Clarke S, & Banfield K. (2001). Homocysteine in health and disease | Hematology. Cambridge University Press. [Google Scholar]
  23. Coppock DL, & Pardee AB (1987). Control of thymidine kinase mRNA during the cell cycle. Molecular and Cellular Biology, 7, 2925–2932. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Crott JW, Liu Z, Choi SW, & Mason JB (2007). Folate depletion in human lymphocytes up-regulates p53 expression despite marked induction of strand breaks in exons 5–8 of the gene. Mutation Research—Genetic Toxicology and Environmental Mutagenesis, 626, 171–179. [DOI] [PubMed] [Google Scholar]
  25. D’Angiolella V, Donato V, Forrester FM, Jeong YT, Pellacani C, Kudo Y, … Pagano M. (2012). Cyclin F-mediated degradation of ribonucleotide reductase M2 controls genome integrity and DNA repair. Cell, 149, 1023–1034. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. D’Angiolella V, Esencay M, & Pagano M. (2013). A cyclin without cyclin-dependent kinases: Cyclin F controls genome stability through ubiquitin-mediated proteolysis. Trends in Cell Biology, 23, 135–140. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. DeGregori J, Kowalik T, & Nevins JR (1995). Cellular targets for activation by the E2F1 transcription factor include DNA synthesis- and G1/S-regulatory genes. Molecular and Cellular Biology, 15, 4215–4224. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Dolnick BJ (2003). Regulation of thymidylate synthase gene expression and drug response. Fluoropyrimidines in Cancer Therapy, 83–91. 10.1007/978-1-59259-337-8_5 [DOI] [Google Scholar]
  29. Doucette MM, & Stevens VL (2001). Folate receptor function is regulated in response to different cellular growth rates in cultured mammalian cells. The Journal of Nutrition, 131, 2819–2825. [DOI] [PubMed] [Google Scholar]
  30. Ducker GS, & Rabinowitz JD (2017). One-carbon metabolism in health and disease. Cell Metabolism, 25, 27–42. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Engström Y, Eriksson S, Jildevik I, Skog S, Thelander L, & Tribukait B. (1985). Cell cycle-dependent expression of mammalian ribonucleotide reductase. Differential regulation of the two subunits. The Journal of Biological Chemistry, 260, 9114–9116. [PubMed] [Google Scholar]
  32. Ercikan-Abali EA, Banerjee D, Waltham MC, Skacel N, Scotto KW, & Bertino JR (1997). Dihydrofolate reductase protein inhibits its own translation by binding to dihydrofolate reductase mRNA sequences within the coding region †. Biochemistry, 36, 12317–12322. [DOI] [PubMed] [Google Scholar]
  33. Eriksson S, Gräslund A, Skog S, Thelander L, & Tribukait B. (1984). Cell cycle-dependent regulation of mammalian ribonucleotide reductase. The S phase-correlated increase in subunit M2 is regulated by de novo protein synthesis. Journal of Biological Chemistry, 259, 11695–11700. [PubMed] [Google Scholar]
  34. Evans DR, & Guy HI (2004). Mammalian pyrimidine biosynthesis: Fresh insights into an ancient pathway. The Journal of Biological Chemistry, 279, 33035–33038. [DOI] [PubMed] [Google Scholar]
  35. Farnham PJ, & Schimke RT (1985). Transcriptional regulation of mouse dihydrofolate reductase in the cell cycle. The Journal of Biological Chemistry, 260, 7675–7680. [PubMed] [Google Scholar]
  36. Feder JN, Assaraf YG, Seamer LC, & Schimke RT (1989). The pattern of dihydrofolate reductase expression through the cell cycle in rodent and human cultured cells. The Journal of Biological Chemistry, 264, 20583–20590. [PubMed] [Google Scholar]
  37. Feder JN, Guidos CJ, Kusler B, Carswell C, Lewis D, & Schimke RT (1990). A cell cycle analysis of growth-related genes expressed during T lymphocyte maturation. The Journal of Cell Biology, 111, 2693–2701. [DOI] [PMC free article] [PubMed] [Google Scholar]
  38. Field MS, Anderson DD, & Stover PJ (2011). Mthfs is an essential gene in mice and a component of the Purinosome. Frontiers in Genetics, 2, 36. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Field MS, Kamynina E, Agunloye OC, Liebenthal RP, Lamarre SG, Brosnan ME, … Stover PJ (2014). Nuclear enrichment of folate cofactors and methylenetetrahydrofolate dehydrogenase 1 (MTHFD1) protect de novo thymidylate biosynthesis during folate deficiency. The Journal of Biological Chemistry, 289, 29642–29650. [DOI] [PMC free article] [PubMed] [Google Scholar]
  40. Field MS, Kamynina E, Watkins D, Rosenblatt DS, & Stover PJ (2015). Human mutations in methylenetetrahydrofolate dehydrogenase 1 impair nuclear de novo thymidylate biosynthesis. Proceedings of the National Academy of Sciences of the United States of America, 112, 400–405. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Field MS, Szebenyi DME, & Stover PJ (2006). Regulation of de novo purine biosynthesis by methenyltetrahydrofolate synthetase in neuroblastoma. The Journal of Biological Chemistry, 281, 4215–4221. [DOI] [PubMed] [Google Scholar]
  42. Fox JT, & Stover PJ (2008). Folate-mediated one-carbon metabolism. Vitamins and Hormones, 79, 1–44. [DOI] [PubMed] [Google Scholar]
  43. Fridman A, Saha A, Chan A, Casteel DE, Pilz RB, & Boss GR (2013). Cell cycle regulation of purine synthesis by phosphoribosyl pyrophosphate and inorganic phosphate. The Biochemical Journal, 454, 91–99. [DOI] [PubMed] [Google Scholar]
  44. Friso S, Choi S-W, Dolnikowski GG, & Selhub J. (2002). A method to assess genomic DNA methylation using high-performance liquid chromatography/electrospray ionization mass spectrometry. Analytical Chemistry, 74, 4526–4531. [DOI] [PubMed] [Google Scholar]
  45. Friso S, Choi SW, Girelli D, Mason JB, Dolnikowski GG, Bagley PJ, … Selhub J. (2002). A common mutation in the 5,10-methylenetetrahydrofolate reductase gene affects genomic DNA methylation through an interaction with folate status. Proceedings of the National Academy of Sciences of the United States of America, 99, 5606–5611. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Girgis S, Suh JR, Jolivet J, & Stover PJ (1997). 5-Formyltetrahydrofolate regulates homocysteine remethylation in human neuroblastoma. The Journal of Biological Chemistry, 272, 4729–4734. [DOI] [PubMed] [Google Scholar]
  47. Good L, Dimri GP, Campisi J, & Chen KY (1996). Regulation of dihydrofolate reductase gene expression and E2F components in human diploid fibroblasts during growth and senescence. Journal of Cellular Physiology, 168, 580–588. [DOI] [PubMed] [Google Scholar]
  48. Grant GD, Brooks L 3rd, Zhang X, Mahoney JM, Martyanov V, Wood TA, … Whitfield ML (2013). Identification of cell cycle-regulated genes periodically expressed in U2OS cells and their regulation by FOXM1 and E2F transcription factors. Molecular Biology of the Cell, 24, 3634–3650. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Graves LM, Guy HI, Kozlowski P, Huang M, Lazarowski E, Pope RM, … Evans DR (2000). Regulation of carbamoyl phosphate synthetase by MAP kinase. Nature, 403, 328–332. [DOI] [PubMed] [Google Scholar]
  50. Gustafsson Sheppard N, Jarl L, Mahadessian D, Strittmatter L, Schmidt A, Madhusudan N, … Nilsson R. (2015). The folate-coupled enzyme MTHFD2 is a nuclear protein and promotes cell proliferation. Scientific Reports, 5, 15029. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Hall CA (1984). The uptake of vitamin B12 by human lymphocytes and the relationships to the cell cycle. The Journal of Laboratory and Clinical Medicine, 103, 70–81. [PubMed] [Google Scholar]
  52. Hendriks IA, D’Souza RC, Chang J-G, Mann M, & Vertegaal ACO (2015). System-wide identification of wild-type SUMO-2 conjugation sites. Nature Communications, 6, 7289. [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Huang C, Sloan EA, & Boerkoel CF (2003). Chromatin remodeling and human disease. Current Opinion in Genetics & Development, 13, 246–252. [DOI] [PubMed] [Google Scholar]
  54. Hurford RK, Cobrinik D, Lee MH, & Dyson N. (1997). pRB and p107/p130 are required for the regulated expression of different sets of E2F responsive genes. Genes & Development, 11, 1447–1463. [DOI] [PubMed] [Google Scholar]
  55. Huynh KM, Kim G, Kim DJ, Yang SJ, Park SM, Yeom YI, … Kang D. (2009). Gene expression analysis of terminal differentiation of human melanoma cells highlights global reductions in cell cycle-associated genes. Gene, 433, 32–39. [DOI] [PubMed] [Google Scholar]
  56. Huynh KM, Soh JW, Dash R, Sarkar D, Fisher PB, & Kang D. (2011). FOXM1 expression mediates growth suppression during terminal differentiation of HO-1 human metastatic melanoma cells. Journal of Cellular Physiology, 226, 194–204. [DOI] [PubMed] [Google Scholar]
  57. Ishida S, Huang E, Zuzan H, Spang R, Leone G, West M, & Nevins JR (2001). Role for E2F in control of both DNA replication and mitotic functions as revealed from DNA microarray analysis. Molecular and Cellular Biology, 21, 4684–4699. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Iwahana H, Honda S, Tsujisawa T, Takahashi Y, Adzuma K, Katashima R, … Itakura M. (1995). Rat genomic structure of amidophosphoribosyltransferase, cDNA sequence of aminoimidazole ribonucleotide carboxylase, and cell cycle-dependent expression of these two physically linked genes. Biochimica et Biophysica Acta, 1261, 369–380. [DOI] [PubMed] [Google Scholar]
  59. Jackson RC, Lui MS, Boritzki TJ, Morris HP, & Weber G. (1980). Purine and pyrimidine nucleotide patterns of normal, differentiating, and regenerating liver and of hepatomas in rats. Cancer Research, 40. [PubMed] [Google Scholar]
  60. Jaenisch R, & Bird A. (2003). Epigenetic regulation of gene expression: How the genome integrates intrinsic and environmental signals. Nature Genetics, 33(Suppl), 245–254. [DOI] [PubMed] [Google Scholar]
  61. Jenh CH, Geyer PK, & Johnson LF (1985). Control of thymidylate synthase mRNA content and gene transcription in an overproducing mouse cell line. Molecular and Cellular Biology, 5, 2527–2532. [DOI] [PMC free article] [PubMed] [Google Scholar]
  62. Johansson E, Hjortsberg K, & Thelander L. (1998). Two YY-1-binding proximal elements regulate the promoter strength of the TATA-less mouse ribonucleotide reductase R1 gene. The Journal of Biological Chemistry, 273, 29816–29821. [DOI] [PubMed] [Google Scholar]
  63. Kaczmarek L. (1986). Protooncogene expression during the cell cycle. Laboratory Investigation, 54, 365–376. [PubMed] [Google Scholar]
  64. Kauffman MG, & Kelly TJ (1991). Cell cycle regulation of thymidine kinase: Residues near the carboxyl terminus are essential for the specific degradation of the enzyme at mitosis. Molecular and Cellular Biology, 11, 2538–2546. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Ke Y, Ash J, & Johnson LF (1996). Splicing signals are required for S-phase regulation of the mouse thymidylate synthase gene. Molecular and Cellular Biology, 16, 376–383. [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Keyomarsi K, Samet J, Molnar G, & Pardee AB (1993). The thymidylate synthase inhibitor, ICI D1694, overcomes translational detainment of the enzyme. The Journal of Biological Chemistry, 268, 15142–15149. [PubMed] [Google Scholar]
  67. Khan S, Abdelrahim M, Samudio I, & Safe S. (2003). Estrogen receptor/Sp1 complexes are required for induction of cad gene expression by 17β-estradiol in breast cancer cells. Endocrinology, 144, 2325–2335. [DOI] [PubMed] [Google Scholar]
  68. Kondo M, Yamaoka T, Honda S, Miwa Y, Katashima R, Moritani M, … Itakura M. (2000). The rate of cell growth is regulated by purine biosynthesis via ATP production and G(1) to S phase transition. Journal of Biochemistry, 128, 57–64. [DOI] [PubMed] [Google Scholar]
  69. Lane AN, & Fan TW-M (2015). Regulation of mammalian nucleotide metabolism and biosynthesis. Nucleic Acids Research, 43, 2466–2485. [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Le François BG, Maroun JA, & Birnboim HC (2007). Expression of thymidylate synthase in human cells is an early G(1) event regulated by CDK4 and p16INK4A but not E2F. British Journal of Cancer, 97, 1242–1250. [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Lee Y, & Johnson LF (2000). Transcriptional control elements of the rat thymidylate synthase promoter: Evolutionary conservation of regulatory features. Experimental Cell Research, 258, 53–64. [DOI] [PubMed] [Google Scholar]
  72. Lewis JA, & Matkovich DA (1986). Genetic determinants of growth phase-dependent and adenovirus 5-responsive expression of the Chinese hamster thymidine kinase gene are contained within thymidine kinase mRNA sequences. Molecular and Cellular Biology, 6(6), 2262–2266. [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Leys EJ, Crouse GF, & Kellems RE (1984). Dihydrofolate reductase gene expression in cultured mouse cells is regulated by transcript stabilization in the nucleus. The Journal of Cell Biology, 99, 180–187. [DOI] [PMC free article] [PubMed] [Google Scholar]
  74. Lin BF, Huang RF, & Shane B. (1993). Regulation of folate and one-carbon metabolism in mammalian cells. III. Role of mitochondrial folylpoly-gamma-glutamate synthetase. Journal of Biological Chemistry, 268, 21674–21679. [PubMed] [Google Scholar]
  75. Littlefield JW (1966). The periodic synthesis of thymidine kinase in mouse fibroblasts. Biochimica et Biophysica Acta, 114, 398–403. [DOI] [PubMed] [Google Scholar]
  76. Liu HT, Baserga R, & Mercer WE (1985). Adenovirus type 2 activates cell cycle-dependent genes that are a subset of those activated by serum. Molecular and Cellular Biology, 5, 2936–2942. [DOI] [PMC free article] [PubMed] [Google Scholar]
  77. Liu HT, Gibson CW, Hirschhorn RR, Rittling S, Baserga R, & Mercer WE (1985). Expression of thymidine kinase and dihydrofolate reductase genes in mammalian ts mutants of the cell cycle. The Journal of Biological Chemistry, 260, 3269–3274. [PubMed] [Google Scholar]
  78. Misselbeck K, Marchetti L, Field MS, Scotti M, Priami C, & Stover PJ (2017). A hybrid stochastic model of folate-mediated one-carbon metabolism: Effect of the common C677T MTHFR variant on de novo thymidylate biosynthesis. Scientific Reports, 7, 797. [DOI] [PMC free article] [PubMed] [Google Scholar]
  79. Mitchell AD, & Hoogenraad NJ (1975). De novo pyrimidine nucleotide biosynthesis in synchronized rat hepatoma (HTC) cells and mouse embryo fibroblast (3T3) cells. Experimental Cell Research, 93, 105–110. [DOI] [PubMed] [Google Scholar]
  80. Morrow GP, MacMillan L, Lamarre SG, Young SK, MacFarlane AJ, Brosnan ME, & Brosnan JT (2015). In vivo kinetics of formate metabolism in folate-deficient and folate-replete rats. The Journal of Biological Chemistry, 290, 2244–2250. [DOI] [PMC free article] [PubMed] [Google Scholar]
  81. Niida H, Katsuno Y, Sengoku M, Shimada M, Yukawa M, Ikura M, … Nakanishi M. (2010). Essential role of Tip60-dependent recruitment of ribonucleotide reductase at DNA damage sites in DNA repair during G1 phase. Genes & Development, 24, 333–338. [DOI] [PMC free article] [PubMed] [Google Scholar]
  82. Nilsson R, Jain M, Madhusudhan N, Sheppard NG, Strittmatter L, Kampf C, … Mootha VK (2014). Metabolic enzyme expression highlights a key role for MTHFD2 and the mitochondrial folate pathway in cancer. Nature Communications, 5, 3128. [DOI] [PMC free article] [PubMed] [Google Scholar]
  83. Noguchi H, Prem veer Reddy G, & Pardee AB (1983). Rapid incorporation of label from ribonucleoside disphosphates into DNA by a cell-free high molecular weight fraction from animal cell nuclei. Cell, 32, 443–451. [DOI] [PubMed] [Google Scholar]
  84. Nordlund P, & Reichard P. (2006). Ribonucleotide reductases. Annual Review of Biochemistry, 75, 681–706. [DOI] [PubMed] [Google Scholar]
  85. Paone A, Marani M, Fiascarelli A, Rinaldo S, Giardina G, Contestabile R, … Cutruzzolà F. (2014). SHMT1 knockdown induces apoptosis in lung cancer cells by causing uracil misincorporation. Cell Death & Disease, 5, e1525. [DOI] [PMC free article] [PubMed] [Google Scholar]
  86. Park K-K, Rue SW, Lee IS, Kim HC, Lee IK, Ahn JD, … Chang YC (2003). Modulation of Sp1-dependent transcription by a cis-acting E2F element in dhfr promoter. Biochemical and Biophysical Research Communications, 306, 239–243. [DOI] [PubMed] [Google Scholar]
  87. Pelka-Fleischer R, Fleischer W, Wilmanns W, Sauer H, & Schalhorn A. (1989). Relation between cell cycle stage and the activity of DNA-synthesizing enzymes in cultured human lymphoblasts: Investigations on cells separated according to DNA content by way of a cell sorter. Leukemia, 3, 380–385. [PubMed] [Google Scholar]
  88. Pelka-Fleischer R, Ruppelt W, Wilmanns W, Sauer H, & Schalhorn A. (1987). Relation between cell cycle stage and the activity of DNA-synthesizing enzymes in cultured human lymphoblasts: Investigations on cell fractions enriched according to cell cycle stages by way of centrifugal elutriation. Leukemia, 1, 182–187. [PubMed] [Google Scholar]
  89. Peña-Diaz J, Hegre SA, Anderssen E, Aas PA, Mjelle R, Gilfillan GD, … Sætrom P. (2013). Transcription profiling during the cell cycle shows that a subset of Polycomb-targeted genes is upregulated during DNA replication. Nucleic Acids Research, 41, 2846–2856. [DOI] [PMC free article] [PubMed] [Google Scholar]
  90. Piper AA, Tattersall MH, & Fox RM (1980). The activities of thymidine metabolising enzymes during the cell cycle of a human lymphocyte cell line LAZ-007 synchronised by centrifugal elutriation. Biochimica et Biophysica Acta, 633, 400–409. [DOI] [PubMed] [Google Scholar]
  91. Polager S, Kalma Y, Berkovich E, & Ginsberg D. (2002). E2Fs up-regulate expression of genes involved in DNA replication, DNA repair and mitosis. Oncogene, 21, 437–446. [DOI] [PubMed] [Google Scholar]
  92. Powell CM, Rudge TL, Zhu Q, Johnson LF, & Hansen U. (2000). Inhibition of the mammalian transcription factor LSF induces S-phase-dependent apoptosis by downregulating thymidylate synthase expression. The EMBO Journal, 19, 4665–4675. [DOI] [PMC free article] [PubMed] [Google Scholar]
  93. Proud CG (2010). The enzymes (Vol. 27, pp. 129–146). [Google Scholar]
  94. Rao GN, & Church RL (1988). Regulation of CAD gene expression in mouse fibroblasts during the transition from the resting to the growing state. Experimental Cell Research, 178, 449–456. [DOI] [PubMed] [Google Scholar]
  95. Rao GN, Church RL, & Davidson JN (1988). Posttranscriptional regulation of the expression of CAD gene during differentiation of F9 teratocarcinoma cells by induction with retinoic acid and dibutyryl cyclic AMP. FEBS Letters, 232, 238–242. [DOI] [PubMed] [Google Scholar]
  96. Rao GN, & Davidson JN (1988). CAD gene expression in serum-starved and serum-stimulated hamster cells. DNA, 7, 423–432. [DOI] [PubMed] [Google Scholar]
  97. Robitaille AM, Christen S, Shimobayashi M, Cornu M, Fava LL, Moes S, … Hall MN (2013). Quantitative phosphoproteomics reveal mTORC1 activates de novo pyrimidine synthesis. Science, 339, 1320–1323. [DOI] [PubMed] [Google Scholar]
  98. Rosario FJ, Powell TL, & Jansson T. (2016). Mechanistic target of rapamycin (mTOR) regulates trophoblast folate uptake by modulating the cell surface expression of FR-α and the RFC. Scientific Reports, 6, 31705. [DOI] [PMC free article] [PubMed] [Google Scholar]
  99. Santiago C, Collins M, & Johnson LF (1984). In vitro and in vivo analysis of the control of dihydrofolate reductase gene transcription in serum-stimulated mouse fibroblasts. Journal of Cellular Physiology, 118, 79–86. [DOI] [PubMed] [Google Scholar]
  100. Schilling LJ, & Farnham PJ (1994). Transcriptional regulation of the dihydrofolate reductase/rep-3 locus. Critical Reviews in Eukaryotic Gene Expression, 4, 19–53. [DOI] [PubMed] [Google Scholar]
  101. Selhub J. (1999). Homocysteine metabolism. Annual Review of Nutrition, 19, 217–246. [DOI] [PubMed] [Google Scholar]
  102. Sherley JL, & Kelly TJ (1988). Regulation of human thymidine kinase during the cell cycle. The Journal of Biological Chemistry, 263, 8350–8358. [PubMed] [Google Scholar]
  103. Shin YS, Chan C, Vidal AJ, Brody T, & Stokstad EL (1976). Subcellular localization of gamma-glutamyl carboxypeptidase and of folates. Biochimica et Biophysica Acta, 444, 794–801. [DOI] [PubMed] [Google Scholar]
  104. Sigoillot FD, Berkowski JA, Sigoillot SM, Kotsis DH, & Guy HI (2003). Cell cycle-dependent regulation of pyrimidine biosynthesis. The Journal of Biological Chemistry, 278, 3403–3409. [DOI] [PubMed] [Google Scholar]
  105. Sigoillot FD, Evans DR, & Guy HI (2002). Growth-dependent regulation of mammalian pyrimidine biosynthesis by the protein kinase A and MAPK signaling cascades. The Journal of Biological Chemistry, 277, 15745–15751. [DOI] [PubMed] [Google Scholar]
  106. Sigoillot FD, Sigoillot SM, & Guy HI (2004). Breakdown of the regulatory control of pyrimidine biosynthesis in human breast cancer cells. International Journal of Cancer, 109, 491–498. [DOI] [PubMed] [Google Scholar]
  107. Slansky JE, & Farnham PJ (1996). Transcriptional regulation of the dihydrofolate reductase gene. BioEssays, 18, 55–62. [DOI] [PubMed] [Google Scholar]
  108. Slansky JE, Li Y, Kaelin WG, & Farnham PJ (1993). A protein synthesis-dependent increase in E2F1 mRNA correlates with growth regulation of the dihydrofolate reductase promoter. Molecular and Cellular Biology, 13, 1610–1618. [DOI] [PMC free article] [PubMed] [Google Scholar]
  109. Smith ML, & Buchanan JM (1979). Nucleotide and pentose synthesis after serum-stimulation of resting 3T6 fibroblasts. Journal of Cellular Physiology, 101, 293–309. [DOI] [PubMed] [Google Scholar]
  110. Solans A, Estivill X, & de la Luna S. (2000). Cloning and characterization of human FTCD on 21q22.3, a candidate gene for glutamate formiminotransferase deficiency. Cytogenetics and Cell Genetics, 88, 43–49. [DOI] [PubMed] [Google Scholar]
  111. Stewart CJ, Ito M, & Conrad SE (1987). Evidence for transcriptional and post-transcriptional control of the cellular thymidine kinase gene. Molecular and Cellular Biology, 7, 1156–1163. [DOI] [PMC free article] [PubMed] [Google Scholar]
  112. Stover P, & Schirch V. (1991). 5-formyltetrahydrofolate polyglutamates are slow tight binding inhibitors of serine hydroxymethyltransferase. The Journal of Biological Chemistry, 266, 1543–1550. [PubMed] [Google Scholar]
  113. Stover PJ, Chen LH, Suh JR, Stover DM, Keyomarsi K, & Shane B. (1997). Molecular cloning, characterization, and regulation of the human mitochondrial serine hydroxymethyltransferase gene. The Journal of Biological Chemistry, 272, 1842–1848. [DOI] [PubMed] [Google Scholar]
  114. Suh JR, Herbig AK, & Stover PJ (2001). New perspectives on folate catabolism. Annual Review of Nutrition, 21, 255–282. [DOI] [PubMed] [Google Scholar]
  115. Suh JR, Oppenheim EW, Girgis S, & Stover PJ (2000). Purification and properties of a folate-catabolizing enzyme. The Journal of Biological Chemistry, 275, 35646–35655. [DOI] [PubMed] [Google Scholar]
  116. Tammsalu T, Matic I, Jaffray EG, Ibrahim AFM, Tatham MH, & Hay RT (2014). Proteome-wide identification of SUMO2 modification sites. Science Signaling, 7, rs2–rs2. [DOI] [PMC free article] [PubMed] [Google Scholar]
  117. Tang J, Liu Z, Ji F, Li Y, Liu J, Song J, … Zhou J. (2015). The role of the cell cycle in the cellular uptake of folate-modified poly(L-amino acid) micelles in a cell population. Nanoscale, 7, 20397–20404. [DOI] [PubMed] [Google Scholar]
  118. Thorndike J, Pelliniemi TT, & Beck WS (1979). Serine hydroxymethyltransferase activity and serine incorporation in leukocytes. Cancer Research, 39, 3435–3440. [PubMed] [Google Scholar]
  119. Tibbetts AS, & Appling DR (2010). Compartmentalization of Mammalian folate-mediated one-carbon metabolism. Annual Review of Nutrition, 30, 57–81. [DOI] [PubMed] [Google Scholar]
  120. Travali S, Lipson KE, Jaskulski D, Lauret E, & Baserga R. (1988). Role of the promoter in the regulation of the thymidine kinase gene. Molecular and Cellular Biology, 8, 1551–1557. [DOI] [PMC free article] [PubMed] [Google Scholar]
  121. Tufi R, Gandhi S, de Castro IP, Lehmann S, Angelova PR, Dinsdale D, … Martins LM (2014). Enhancing nucleotide metabolism protects against mitochondrial dysfunction and neurodegeneration in a PINK1 model of Parkinson’s disease. Nature Cell Biology, 16, 157–166. [DOI] [PMC free article] [PubMed] [Google Scholar]
  122. Whitfield ML, Sherlock G, Saldanha AJ, Murray JI, Ball CA, Alexander KE, … Botstein D. (2002). Identification of genes periodically expressed in the human cell cycle and their expression in tumors. Molecular Biology of the Cell, 13, 1977–2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  123. Woeller CF, Anderson DD, Szebenyi DME, & Stover PJ (2007). Evidence for small ubiquitin-like modifier-dependent nuclear import of the thymidylate biosynthesis pathway. The Journal of Biological Chemistry, 282, 17623–17631. [DOI] [PubMed] [Google Scholar]
  124. Wu J-SR, & Johnson LF (1982). Regulation of dihydrofolate reductase gene transcription in methotrexate-resistant mouse fibroblasts. Journal of Cellular Physiology, 110, 183–189. [DOI] [PubMed] [Google Scholar]
  125. Yamaoka T, Kondo M, Honda S, Iwahana H, Moritani M, Ii S, … Itakura M. (1997). Amidophosphoribosyltransferase limits the rate of cell growth-linked de novo purine biosynthesis in the presence of constant capacity of salvage purine biosynthesis. The Journal of Biological Chemistry, 272, 17719–17725. [DOI] [PubMed] [Google Scholar]
  126. Zhu B, Xiahou Z, Zhao H, Peng B, Zhao H, & Xu X. (2014). MTHFR promotes heterochromatin maintenance. Biochemical and Biophysical Research Communications, 447, 702–706. [DOI] [PubMed] [Google Scholar]

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