Accepted Manuscript Pyrimidine Metabolism: Dynamic and Versatile Pathways in Pathogens and Cellular Development Manuel F. Garavito, Heidy Y. Narváez-Ortiz, Barbara H. Zimmermann PII:

S1673-8527(15)00076-4

DOI:

10.1016/j.jgg.2015.04.004

Reference:

JGG 363

To appear in:

Journal of Genetics and Genomics

Received Date: 21 November 2014 Revised Date:

13 April 2015

Accepted Date: 14 April 2015

Please cite this article as: Garavito, M.F., Narváez-Ortiz, H.Y., Zimmermann, B.H., Pyrimidine Metabolism: Dynamic and Versatile Pathways in Pathogens and Cellular Development, Journal of Genetics and Genomics (2015), doi: 10.1016/j.jgg.2015.04.004. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Pyrimidine Metabolism: Dynamic and Versatile Pathways in Pathogens and Cellular

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Development

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Manuel F. Garavito1, Heidy Y. Narváez-Ortiz1 and Barbara H. Zimmermann*

Departamento de Ciencias Biologicas, Universidad de los Andes, Carrera 1, No. 18A-10,

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Bogotá, Colombia

*Corresponding author. Tel.: 571 339 4949 ext 3374; fax: 571 613 0222. [email protected]

authors contributed equally to this work.

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‡These

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Abbreviations: CAD, multifunctional protein that initiates and regulates de novo pyrimidine biosynthesis; DHODase, dihydroorotate dehydrogenase; hpi, hours postinfection; UMP synthase, uridine 5′-monophosphate synthase; UMP, uridine 5′-monophosphate; UTP, uridine 5′triphosphate; UPRTase, uracil phosphoribosyltransferase; UPase, uridine phosphorylase.

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Abstract

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The importance of pyrimidines lies in the fact that they are structural components of a broad

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spectrum of key molecules that participate in diverse cellular functions, such as synthesis of

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DNA, RNA, lipids, and carbohydrates.

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involved in the synthesis, degradation, salvage and interconversion and transport of these

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molecules.

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metabolism changes under a variety of conditions including, when possible, those studies based

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on techniques of genomics, transcriptomics, proteomics, and metabolomics. First, we briefly

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look at the dynamics of pyrimidine metabolism during nonpathogenic cellular events. We then

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focus on changes that pathogen infections cause in the pyrimidine metabolism of their host.

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Next, we discuss the effects of antimetabolites and inhibitors, and finally we consider the

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consequences of genetic manipulations, such as knock-downs, knock-outs, and knock-ins, of

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pyrimidine enzymes on pyrimidine metabolism in the cell.

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Keywords:

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dehydrogenase, UMP synthase.

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(1) Introduction.

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Pyrimidines are essential biomolecules. They are structural components of nucleotides, nucleic

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acids, vitamins, pterins and folates, each fulfilling critical roles in the cell (Cox, 1968). The

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metabolism of pyrimidines has been broadly studied in many organisms. Their critical

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participation in processes such as RNA and DNA synthesis, formation of UDP-sugars for

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glycosylation of proteins and lipids, and formation of CDP-activated precursors for membrane

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phospholipids, makes them attractive for study.

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Pyrimidine metabolism encompasses all enzymes

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In this review, we discuss recent publications that document how pyrimidine

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pyrimidine metabolism, pathogens, CAD, dihydroorotase, dihydroorotate

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Pyrimidine derivatives have been used as anticancer, antimicrobial, antiviral, antibacterial,

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antiprotozoal, and antifungal agents. Pharmacological applications of pyrimidine-related

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molecules are even more diverse, ranging from antipyretics, antihypertensives, antihistaminics,

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anti-inflammatories, analgesics, antidiabetics, antiallergics, antioxidants, anticonvulsants, to

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central nervous system depressants (Jain et al., 2006; Chauhan and Kumar, 2013; Sharma et al.,

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2014). The study of pyrimidine metabolism is important for better understanding of both the

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biochemistry of the use of these compounds, as well as the metabolism of drugs derived from

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pyrimidines. Fig. 1 shows most pathways of pyrimidine metabolism. There is a consensus on what

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constitutes the de novo and degradation pathways, however, the subdivision of pyrimidine

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metabolism reactions into salvage or interconversion pathways is controversial. We have chosen

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to group the reactions according to whether they involve nucleobases, nucleosides, nucleosides

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mono-, di- or triphosphates (nucleotides).

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The de novo pathway is initiated with bicarbonate, glutamine and two molecules of ATP.

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This pathway involves six enzymatic steps conserved in all species, although they have different

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architecture depending on the organism (for detailed information see Nara et al., 2000). In most

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prokaryotes and some simple eukaryotes, the enzymes of the de novo pathway are encoded by

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separate genes, whereas in mammals they are encoded by only three genes, carbamoylphosphate

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synthetase–aspartate transcarbamoylase–dihydroorotase (CAD), dihydroorotate dehydrogenase

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(DHODase) and uridine 5′-monophosphate synthase (UMP synthase), (Jones, 1980). The

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trifunctional enzyme CAD converts the small precursors into dihydroorotic acid that is

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transformed into orotate by DHODase. Subsequently, the bifunctional enzyme UMP synthase

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leads to the synthesis of uridine 5′-monophosphate (UMP). UMP could be considered one of the

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hub molecules in pyrimidine metabolism because it is the precursor of other pyrimidine

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nucleotides.

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The nucleoside di- and triphosphates are formed by nucleoside kinases, and nucleoside

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diphosphates can be converted into the deoxynucleotides by the ribonucleotide reductase

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complex. At the bottom of Fig. 1, we show reactions involving nucleotides that are necessary for

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forming larger biomolecules such as nucleic acids, lipids and carbohydrates (for detailed

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information see Huang and Graves, 2003).

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Membrane transporters also play important roles in pyrimidine metabolism as shown at the

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top of Fig. 1. In humans, transporters are needed for importing amino acids, nucleobases and

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nucleosides. The solute carrier (SLC) transporter family 17 proteins permit the exchange of

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amino acids essential for pyrimidine biosynthesis such as aspartate and glutamate, and

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nucleotides (Sreedharan et al., 2010). Equilibrative nucleoside transporters (ENTs) are

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responsible for transport of hydrophilic nucleosides. Concentrative nucleoside transporters

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(CNT) transport nucleobases across membranes, and transporters from the ABC transporter

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family import nucleotides and nucleosides into the cell (Sahoo et al., 2014). Pyrimidines can be degraded to simple metabolites, which become sources of carbon and

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nitrogen. While in some organisms such as Escherichia coli, there are multiple catabolic

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pathways, nucleosides are catabolized mainly by two pathways, one oxidative the other reductive

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(Loh et al., 2006). The oxidative pathway (not shown in Fig.1) has been reported only in some

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bacteria and produces urea as a primary product, and malonic acid from uridine/cytidine, or

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methylmalonic acid from thymine, as secondary products (Patel and West, 1987). The reductive

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pathway (Fig. 1, right), which is present in some archaea, bacteria and eukaryotes, consists of

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three catalytic steps to produce CO2, NH3 and ß-alanine (ß-Ala) or ß-aminoisobutyric acid (ß-

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AIB). The nucleosides, uridine and thymidine can be reductively catabolized when they are

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transformed to nucleobases by the action of the nucleoside phosphorylase enzymes. However,

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cytidine must be deaminated to uridine by cytidine deaminase before entering the catabolic route

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(Vogels and van der Drift, 1976). It has been proposed that the pyrimidine catabolic pathways

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are important not only to maintain appropriate pyrimidine pools in cells, but also to produce ß-

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alanine, a precursor of diverse biomolecules, for example, pantothenic acid, vitamin B5 in plants

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and bacteria (Webb et al., 2004; Katahira and Ashihara, 2006) and a neurotransmitter in

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mammals (Tiedje et al., 2010).

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Different organisms contain different subsets of enzymes in the pyrimidine pathways

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briefly described above. Table S1 indicates which genes of pyrimidine metabolic proteins are

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present in the organisms mentioned in this review. The presence or absence of specific

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pyrimidine enzymes in different species probably evolved depending on the availability and

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accessibility of nutrients in the niche in which each organism developed. It is precisely these

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marked differences between species that makes pyrimidine metabolism a potential drug target.

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For example, protozoan parasites of great medical importance such as Plasmodium, Toxoplasma

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and Cryptosporidium show a reduction in the numbers of genes involved in pyrimidine

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metabolism. Plasmodium parasites are unable to obtain pyrimidines by the salvage pathway and

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depend completely on the de novo pathway (Fig. 2), whereas in Cryptosporidium the opposite

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occurs (Hyde, 2007).

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Our goal in this review is to assemble recent advances pertinent to an overview of

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pyrimidine metabolism. The demand for pyrimidines varies with the overall metabolic activity of

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the cell. It has long been known that actively dividing cells require higher levels of pyrimidines

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than quiescent cells, thus, cancer cells are observed to overexpress pyrimidine metabolic

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enzymes (Evans and Guy, 2004; Hu et al., 2013). Consideration of this last area is beyond the

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scope of this review, and instead we will focus primarily, but not exclusively, on the changes in

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pyrimidine metabolism during the life cycles of some pathogens, including those parasitic stages

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that occur within host organism.

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Some of the first insights into the roles of pathways were gained by measuring enzyme

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activities and by observing the effects of inhibitors.

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permitted monitoring expression levels of enzymes in these pathways. The newest contributions

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to the area of pyrimidine metabolism have been through metabolomics.

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More recently, transcriptomics has

For a very few

organisms, data from all of these areas are available. The integration of genomic, transcriptomic,

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proteomic, and metabolomic data will allow for a better understanding of the dynamics of

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pyrimidine metabolism.

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A commonly-used approach that highlights the importance of specific enzymes in a

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pathway is to interfere with their normal function by using inhibitors. Such studies are critical

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for identifying potential drug targets for pathogens. A more recent strategy that delineates the

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importance of pathway enzymes is to construct genetic knock-outs, knock-ins, or knock-downs.

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Although genetically altered organisms in many cases may have limited relevance for medical

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applications, a study of their metabolism demonstrates the direct effects of increased or

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decreased enzyme activities that are not plagued by unintended secondary effects or poor cell

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permeability associated with inhibitory compounds.

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(2) Changes in pyrimidine metabolism during cellular development.

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Using serum starvation to synchronize a baby hamster kidney cell line and incorporation of

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pyrimidine biosynthesis in the S phase of the cell cycle was ~ 2-fold higher than in other phases

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of the cycle (Sigoillot et al., 2003). The increase in biosynthesis could be accounted for by

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allosteric effects on the carbamoylphosphate synthetase (CPSase) activity of CAD, which

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included removal of feedback inhibition by UTP and increased sensitivity to the activator

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phosphoribosyl pyrophosphate (PRPP), and was mediated by phosphorylation of CAD at Thr-

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456 by MAP kinase (MAPK).

C-labeled bicarbonate into UTP and CTP, Sigoillot and co-workers found that the rate of

As cell cultures became confluent, Thr-456 became

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dephosphorylated, and cAMP-dependent kinase A (PKA) phosphorylated Ser-1406, leading to

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decreased PRPP sensitivity, and consequently decreased CAD CPSase activity. The authors

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measured intracellular nucleotide concentrations and observed that the levels remained

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approximately constant during the cell cycle, and were about half the concentrations found in

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exponentially growing cells. This led to the conclusion that newly synthesized nucleotides are

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used as fast as they are produced (Sigoillot et al., 2003), suggesting that the pyrimidine pools in

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these cells are maintained at subsistence levels. Two reports in 2013 (Ben-Sahra et al., 2013;

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Robitaille et al., 2013) indicated that in addition to the allosteric regulation via the MAPK

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cascade (Erk1/2) signaling pathway, CAD is also subject to allosteric regulation via the

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mTORC1 signaling network which controls S6 kinase (S6K) phosphorylation of CAD at residue

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Ser-1859.

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The yeast Candida albicans is an opportunistic pathogen that causes fungal infections in

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immunocompromised patients.

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electrophoresis to measure the proteins present in stationary and exponential growth phases of

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the pathogen. The stationary phase is probably most relevant to infections because growth is

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constrained by the nutrient-limited environment of the host cell. Five enzymes of pyrimidine

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metabolism were among those identified.

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orotidine 5′-monophosphate decarboxylase (ODCase), and uracil phosphoribosyltransferase

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(UPRTase) showed increased levels in the exponential phase, and cytidine triphosphate synthase

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(CTP synthase) showed higher levels in the stationary phase, while dihydroorotase (DHOase)

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levels were the same in both phases (Kusch et al., 2008). Keeping in mind the caveat that higher

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protein levels do not prove that there is increased flux through a pathway, one could explain the

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observed changes in protein levels as follows. An increase in levels of proteins involved in

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synthesis or recycling of nucleotides might be expected because it could allow the yeast to

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maintain its supply of nucleotides permitting the rapid growth rate of the exponential phase. The

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observation of the same protein levels for the DHOase in both phases may simply mean that it

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does not play a rate-limiting role in the de novo pathway in this organism. The reason for higher

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requirements of CTP synthase in the stationary phase is not clear, and may suggest that this

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protein plays additional roles in the physiology of that phase. This is not surprising because it

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has been recently documented that CTP synthase could also have a structural role in the cell (for

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detailed information see Chen et al., 2011; Liu, 2011; Barry et al., 2014; Carcamo et al., 2014).

Kusch and co-workers (Kusch et al., 2008) used 2D-gel

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Orotate phosphoribosyltransferase (OPRTase),

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Cell death can be viewed as the expected final event in cell development. For this reason,

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understanding the changes in the metabolic environment that lead to and occur during

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programmed cell death (PCD) is important. As might be expected, changes are observed in

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pyrimidine metabolism during PCD. Stasolla and co-workers (Stasolla et al., 2004) incubated

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plant cell cultures with radiolabeled uracil, uridine and thymidine, and measured their

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incorporation into nucleotides and nucleic acids. Degradation was also monitored. PCD was

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induced by exposing the cultures to agents generating NO and H2O2. An early event, occurring

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before cytological hallmarks of PCD were evident, was the reduced incorporation of uracil into

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nucleotides and nucleic acids. Later events of PCD included reduced incorporation of uridine,

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and continued reduced incorporation of uracil, and increased degradation of both compounds. In

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contrast, thymidine incorporation increased (Stasolla et al., 2004). The changes in the balance of

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salvage and degradation of pyrimidines were proposed to constitute a metabolic switch that

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eventually commits the cell to PCD (Stasolla et al., 2004).

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(3) Changes in pyrimidine metabolism during pathogen infection of host cells. To ensure their own survival, many pathogens redirect the metabolism of the cells they

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invade. In this section we will consider the changes in host pyrimidine metabolism observed in

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two host/pathogen systems, first, fibroblasts infected by human cytomegalovirus (HCMV), and

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second, red blood cells infected by Plasmodium falciparum. Another case we will consider is

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that of Phytophthora infestans infecting Solanum tuberosum (potato), which is an example of a

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pathogen modifying expression of its own pyrimidine metabolic enzymes during its life cycle.

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This last point is also illustrated in Fig. 2, which assembles RNA Sequencing (RNA-seq) data of

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pyrimidine metabolic enzymes for seven pathogens.

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Munger and co-workers measured metabolites during the infection of HCMV in quiescent

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fibroblasts (Munger et al., 2006). The time course of the infection was marked by the following

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events: viral gene expression starting at 2 hours postinfection (hpi), viral DNA replication

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between 24 – 30 hpi, peak virus yielding between 72 – 96 hpi, and host cell death. Changes in

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metabolites of infected cells relative to mock-infected cells were observed for the de novo

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pyrimidine biosynthetic pathway. Carbamoyl phosphate, the product of the first step catalyzed by

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CAD, began decreasing at 72 hpi, and at 96 hpi it was ~ 4-fold lower than mock-infected cells

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(see Fig. 4 in Munger et al., 2006). The largest concentration change occurred with carbamoyl

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aspartate, the product of the second CAD reaction, which began to increase at 24 hpi, and

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achieved levels ~ 25-fold higher than mock-infected cells at 80 hpi (see Fig. 4 in Munger et al.,

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2006).

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aforementioned measurements were performed with infected, serum-starved, confluent cells,

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nevertheless, the authors found that these metabolite levels were still substantially higher than

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those of actively growing fibroblasts.

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significantly during infection (Munger et al., 2006). The accumulation of carbamoyl aspartate

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might be explained as follows: the DHOase reaction is known to be reversible (Christopherson

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and Jones, 1979), thus one could speculate that a decoupling of CAD reactions from the reaction

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of the fourth enzyme, DHODase, could be caused by a change in localization of CAD, for

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example from the cytosol to the nucleus (Angeletti and Engler, 1998; Sigoillot et al., 2003), and

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might lead to reversal of the DHOase reaction. In a later publication from the same laboratory, a

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more complete survey of metabolites was made in cells infected by HCMV and also cells

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infected by herpes simplex virus type 1 (HSV-1) (Vastag et al., 2011). The authors concluded

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that the two viruses employ different strategies to promote viral replication. HCMV drives the

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host cell from quiescence toward G1/S, and the production of nucleotides, at the same time

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preventing cellular DNA replication, while HSV-1 encodes necessary enzymes, for example

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thymidine kinase, dUTPase, and uracil-DNA glycosylase, and increases flux through central

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carbon metabolism to drive nucleotide synthesis (Vastag et al., 2011).

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UMP levels decreased slightly, while UTP, CTP, and TTP levels increased.

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Interestingly, purine pool levels did not change

The first study of the metabolome in the red blood cell during the 48 h infection cycle of P.

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falciparum revealed interesting changes in pyrimidine levels (Olszewski et al., 2009).

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Intracellular concentrations of UMP and CMP, increased to peak concentrations of ~ 28-fold and

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~ 24-fold, respectively, relative to uninfected cell at 32 – 48 h (see Table S1 in Olszewski et al.,

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2009). Intracellular concentrations of the de novo synthesis intermediates dihydroorotate and

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orotate followed a similar periodic fluctuation during the cycle, but with much lower increases (~

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4-fold and ~ 2-fold, respectively) (see Table S1 in Olszewski et al., 2009). Concentrations in

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extracellular media were also measured, and showed secretion of ~ 5-fold more dihydroorotate

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from infected cells compared to uninfected cells, peaking at 32 h (see Table S2 in Olszewski et

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al., 2009). A more dramatic increase was observed for deoxyuridine, whose secretion steadily

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increased starting at 24 h and reached ~ 24-fold at 48 h compared to uninfected cells (see Table

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S2 in Olszewski et al., 2009). To better understand the fluctuations in metabolites, the authors

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measured mRNA transcript abundance during infection using DNA microarrays. Surprisingly,

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they found a significant negative correlation between the expression levels of the enzyme

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DHOase and the levels of its product dihydroorotate, that is, enzyme expression levels were

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higher during initial stages of infection, when its product concentrations were low. Later in

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infection, enzyme expression decreased, but product increased.

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apparently contradictory data may lie in analysis of the DHOase mRNA half-life (Shock et al.,

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2007). The biological database for the genus Plasmodium (PlasmoDB) shows that its stability is

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low during the first 24 h, accordingly, mRNA expression must be high to ensure the

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establishment of infection, however, after schizogony (24 h), DHOase mRNA stability is

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enhanced by ~ 8-fold. Thus, the parasite regulates not only mRNA expression but also the

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mRNA stability to ensure the availability of nucleotides.

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The explanation for these

The life cycle of the plant pathogen P. infestans is hemibiotrophic. In the initial stages of

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infection, the biotrophic phase, this oomycete proliferates rapidly without causing visible

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symptoms. The later stages, known as the necrotrophic phase, lead to the death of the plant cell.

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This change of life style is reflected in large shifts in gene expression. A quantitative reverse

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transcription-PCR (qRT-PCR) study of relative expression profiles showed up-regulation of the

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de novo synthetic enzymes DHOase and UMP synthase, accompanied by a concomitant down-

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regulation of the salvage enzyme UPRTase during the biotrophic phase (García-Bayona et al.,

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2014). During the necrotrophic phase, the relative expression showed an inverse behavior,

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apparently relying more on salvage than on synthesis. The authors state that the changes in

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expression could be related to higher levels of uracil, uridine, cytosine, or cytidine that may be

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available from the dying plant cell, or could even be a strategy of the cell leading to an

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imbalance of the nucleotide pools and host cell death phase (García-Bayona et al., 2014).

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In Fig. 2, we have assembled transcription information for the human pathogens

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Toxoplasma gondii (toxoplasmosis), P. falciparum (malaria), Cryptosporidium parvum

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(cryptosporidiosis), Trypanosoma brucei (African trypanosomiasis), and Cryptococcus

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neoformans (meningitis), and for the plant pathogens Colletotrichum higginsianum (anthracnose

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disease) and Phytophthora ramorum (sudden oak death). Parasitic pathogens obtain metabolic

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resources from the host cells they infect, and thus do not code for a full complement of

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pyrimidine metabolic machinery in their genomes. Genes not present are indicated by the color

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gray in Fig. 2. Notably, none of these pathogens have genes for pyrimidine degradation. On the

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far left side of the heat map, we have included data comparing mouse cells infected with T.

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gondii to uninfected mouse cells. Interestingly, infection with T. gondii results in a dramatic

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increase in uridine phosphorylase (UPase), and dihydroopyrimidine dehydrogenase in the mouse

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cells. Comparing transcriptomes of different species using RNA-seq is challenging. One critical

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aspect is that analyses are based on sequence conservation, yet in many cases there is a lack of

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high-quality annotation of orthologous genes. Another difficulty in constructing this type of

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figure was in choosing which two stages or conditions of the organism to compare, and these

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choices were also limited by the availability of data. Additionally, it should be remembered that

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transcriptional up- and down-regulation is just one of many mechanisms used to regulate enzyme

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activities. Bearing these limitations in mind, what is striking in Fig. 2, is the variability in

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transcription patterns observed for these organisms, which no doubt reflect the specialized niches

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that they occupy.

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(4) Effects of antimetabolites and inhibitors on pyrimidine metabolism. An extensive study of the effect of halogenated pyrimidines was undertaken on the

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bloodstream form of the kinetoplastid parasite T. brucei using metabolomics (Ali et al., 2013).

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Transport assays revealed that the only pyrimidine transporter in this form of the parasite was a

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high affinity uracil transporter, also capable of transporting orotate, uridine, and deoxyuridine,

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but with lower affinities. The transporter exhibited 5-fold less affinity for 5-fluorouracil (5-FU)

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than uracil, and the former was salvaged by the parasite’s UPRTase, phosphorylated, and

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incorporated into RNA. Since no incorporation into DNA was observed, 5-FU was not a

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substrate for parasite ribonucleotide reductase, although it is a substrate for human

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ribonucleotide reductase. Incorporation into the UDP-activated hexose and hexosamines was

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observed, although no differences in content of variant surface glycoprotein was found for

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treated parasites, suggesting that 5-FU did not interfere with the parasite’s carbohydrate

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metabolism. 5-fluoroorotate (5-FOA) was metabolized in a similar fashion to 5-FU.

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Unexpectedly, fluoro-N-carbamoyl-L-aspartate was also detected in 5-FOA-treated parasites.

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The authors suggested that this partial reversal of synthesis may have been caused by increased

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levels of orotate, which they attributed to the inhibition of OPRTase by 5-FOA. They noted that

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both compounds caused growth arrest, but did not kill the parasites quickly, and suggested that

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their mechanism of action were probably multifactorial, and due in part to incorporation of

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fluorinated nucleotides into RNA. Parasites treated with 5-fluoro-2′-deoxyuridine produced low

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amounts of 5F-dUMP with thymidine kinase, however, no F-dUDP or F-dUTP could be

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detected, and no incorporation into nucleic acids was observed. Treated parasites exhibited

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elevated levels of dUMP, but had close to normal levels of thymidine nucleotides, apparently due

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to salvage of exogenous thymidine present in the medium, supporting the idea that the effect of

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5-fluoro-2′-deoxyuridine was likely to be caused by inhibition of thymidylate synthase.

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The effect of increased levels of a pyrimidine intermediate may have different consequences

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depending on cell type and conditions. Examples of this are the divergent effects of an inhibitor

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on multiple myeloma cells (Bardeleben et al., 2013) and on hepatocellular carcinoma cells (Jung

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et al., 2012).

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carboxamide-1- β -riboside (AICAr), which becomes converted to AICA ribotide 5′‐

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monophosphate (ZMP), a mimic of AMP, was found to induce apoptosis, however, this was not

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associated with AMP-activated kinase activation or mTORC1 inhibition. A metabolomic screen

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of AICAr-treated cells showed a ~ 26-fold increase in orotate and a ~ 0.4-fold decrease in UMP

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that was accompanied by a 1.2 – 1.9-fold increase in purine bases and nucleotides (see Table 1 in

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Bardeleben et al., 2013). Additionally, a ~ 3-fold decrease in PRPP levels was detected by an

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enzymatic assay. The treated cells were found to be under replicative stress as indicated by the

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phosphorylation of the histone H2A.X, which replaces conventional H2A in a subset of

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nucleosomes. The authors suggested that the orotate accumulation was due to a decrease in

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UMP synthase activity caused by limiting PRPP levels, and found that the treated cells could be

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rescued by exogenous uridine, cytidine, and thymidine. Application of N-(phosphonacetyl)-L-

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aspartate (PALA), a specific transition state analog inhibitor of aspartate transcarbamoylase

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(ATCase), was also found to result in H2A.X phosphorylation, and could be rescued in a similar

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fashion by uridine and cytidine. The authors concluded that apoptosis in AICAr-treated cells

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was caused by pyrimidine starvation (Bardeleben et al., 2013). In contrast, in hepatocarcinoma

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cells, exogenous application of orotate to serum-starved cells induced proliferation by increasing

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the number of viable cells and decreasing apoptosis as mediated by mTORC1 activation (Jung et

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al., 2012). The fate of the orotate after it was internalized by the cells was not determined (Jung

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et al., 2012). These researchers used AICAr to activate AMP-activated protein kinase (AMPK)

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producing negative regulation of mTOR.

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hepatocarcinoma cells with orotate and AICAr resulted in suppression of proliferation (Jung et

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In multiple myeloma cell cultures, the application of 5-aminoimidazole-4-

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al., 2012). In these two studies, AICAr appeared to affect the mTOR pathway differently,

308

however, in both cases the presence of AICAr combined with high levels of orotate, either

309

exogenous or endogenous, led to decrease in proliferation or apoptosis. In human hepatoma cell lines, inhibition of UMP synthase was shown to have potent

311

effects on cholesterol and lipid metabolism (Poirier et al., 2014). The unmethylable cytosine

312

analog 5-azacytidine (5-AzaC) is used as an epigenetic drug which permits cells to re-express

313

genes silenced by hypermethylation. Poirer and co-workers found that 5-AzaC had methylation-

314

independent effects that included decreasing cellular UTP and CTP levels, decreasing mRNA

315

levels of genes involved in cholesterol and lipid metabolism, and increasing triacylglycerol

316

synthesis and cytosolic lipid droplet formation. Notably, these effects could be reversed by

317

adding either UTP or cytidine. Treatment of cells with pyrazofurin, a known inhibitor of UMP

318

synthase, also increased UTP and CTP levels and promoted lipid droplet formation. Although

319

the mechanisms behind the observed changes in lipid and cholesterol metabolism remain to be

320

elucidated, the cross-talk between pyrimidine biosynthesis and glycerolipid has been clearly

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demonstrated.

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(5) Effects of genetic modifications on pyrimidine metabolism.

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The pathogenic fungus C. neoformans is responsible for fatal meningitis in

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immunocompromised patients. A knockout of DHOase (URA4) conferred a growth defect at

326

high temperature, impairing its ability to grow at the physiological temperature of 37°C. RT-

327

PCR revealed that wild type C. neoformans increased expression of genes in both pyrimidine

328

biosynthetic and salvage pathways at restrictive temperatures, in minimal media in the absence

329

of uracil. Under the same conditions, the DHOase knockout mutant exhibited a decrease in

330

expression in biosynthetic genes, while expression of salvage genes remained unchanged. The

331

mutant caused non-lethal infections in mice, nevertheless, the pathogen could be isolated from

332

tissue 40 days post infection (de Gontijo et al., 2014). In a study that sought to explain the

333

synergic effect of 5-fluorocytosine and Amphotericin B combination therapy for cryptococcosis

334

in human immunodeficiency virus (HIV) patients, a C. neoformans knockout of OPRTase

335

(URA5) was constructed. The mutants were found to be hypersensitive to Amphotericin B,

336

probably as a result of a deficiency in the pools of UDP-sugars required for synthesis of the

337

pathogen’s polysaccharide capsule (Banerjee et al., 2014).

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Zameitat and co-workers (2007) deleted the pyr4 gene coding for the mitochondrial

339

DHODase from the basidiomycetous plant pathogen Ustilago maydis. The mutated fungus was

340

auxotrophic for pyrimidines, and exhibited enhanced sensitivity to UV radiation that had been

341

previously reported for mutants deficient in other enzymes of pyrimidine biosynthesis (for

342

example, pyr1). The radiation sensitivity of the pyr4 mutant was completely alleviated by

343

complementation with a human DHODase construct, indicating that the sensitivity is directly

344

related to impairment of de novo synthesis. Importantly, the pyr4 mutant was nonpathogenic in

345

maize, thus de novo synthesis appears to be essential for virulence.

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In the nematode Caenorhabditis elegans, the rad-6 gene, whose mutation results in

347

hypersensitivity to DNA damaging agents, was recently identified as UMP synthase. The

348

hypersensitivity was attributed to the predicted lower concentrations of pyrimidine pools that

349

would decrease their availability for DNA repair. Additionally, mutants exhibited phenotypes

350

associated with defects of proteoglycan synthesis and orotate accumulation, and had reduced

351

lifespans (Merry et al., 2014). When the salvage enzyme uridine kinase (UKase) was knocked

352

down with RNAi in the rad-6 mutant, viability was drastically reduced. In contrast, a UKase

353

knockout in the wild type worm had little effect, suggesting that C. elegans relies more on the de

354

novo pathway than on pyrimidine salvage.

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A similar situation was found in the apicomplexan parasite T. gondii, where mutants

356

lacking the salvage enzymes UPRTase (Pfefferkorn and Pfefferkorn, 1979) or UPase (Fox and

357

Bzik, 2010) were both replication competent and virulent. In contrast, parasites with knockouts

358

of the T. gondii biosynthetic enzymes CPSase (Fox and Bzik, 2002), or ODCase (Fox and Bzik,

359

2010) were avirulent in mice, and were unable to replicate in cell culture. The growth of both of

360

these knockouts could be rescued in vitro with uracil supplementation. Interestingly, it was not

361

possible to construct a DHODase knockout, even in the presence of uracil supplementation

362

(Hortua Triana et al., 2012). In P. falciparum, transgenic parasites expressing a cytosolic,

363

fumarate-dependent yeast DHODase were found to be resistant to specific inhibitors for the

364

endogenous DHODase, and also exhibited resistance to mitochondrial electron transport chain

365

inhibitors (Painter et al., 2007; Ke et al., 2011), nevertheless, a PfDHODase knockout has not

366

been reported. All of the transgenic strains were able to maintain long-term growth in presence

367

of PfDHODase inhibitors, but some of the strains required decylubiquinone supplementation for

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long-term survival in the presence of mitochondrial electron transport chain inhibitors (Ke et al.,

369

2011). In some organisms, genetic manipulations of pyrimidine metabolism lead to the use of

371

alternative pathways. An example is the double knockout of the two alleles of UMP synthase in

372

T. brucei (Ong et al., 2013). While the original mutant parasites were nonvirulent in mice,

373

mutants maintained in long-term tissue culture unexpectedly regained virulence in mice. Uptake

374

studies showed that both original and long-term mutants increased their transport of uracil from

375

the media compared to wild type parasites. Unlike wild type parasites, both original and long-

376

term mutants secreted orotate when grown in pyrimidine-poor environments, however, the

377

latter’s secretion levels were higher. The authors hypothesized that the mutants’ secretion of

378

orotate into the mouse would result in the mouse metabolizing this intermediate into a

379

pyrimidine that the parasite could then transport to supplement its pyrimidine deficiency (Ong et

380

al., 2013).

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Another example of a microorganism exhibiting adaptability in pyrimidine metabolism is

382

Lachancea kluyveri.

383

pyrimidines:

384

dehydrogenase, which is absent in all fungi), and the newly discovered uracil catabolic pathway

385

(URC) pathway (Andersen et al., 2008). Microarray expression analysis was performed to

386

determine which gene transcripts were affected when different pyrimidine metabolites were used

387

as the sole nitrogen source (Andersson Rasmussen et al., 2014). The URC genes were induced

388

by uracil, while reductive pathway genes (PYD) were induced by dihydrouracil. A new gene

389

coding for the enzyme catalyzing the final step in the URC pathway was identified. In addition,

390

eight new genes encoding putative permeases (transporters) were identified. These permeases

391

appeared to exhibit functional redundancy, since all single and double knockouts strains were

392

found to be viable (Andersson Rasmussen et al., 2014).

This yeast possesses two of the four known catabolic pathways for degradation

(excluding

the

first

enzyme,

dihydropyrimidine

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reductive

393

Unexpectedly, antisense knockdown of the UMP synthase of S. tuberosum resulted in

394

increased biosynthesis of starch and cell wall in growing tubers (Geigenberger et al., 2005).

395

Transgenic plants were generated by using antisense inhibition of UMP synthase under the

396

control of a tuber-specific promoter. Plant lines with 30 – 85% reduced enzymatic activity in

397

growing tubers were selected. A decrease in UMP synthase activity of ≥ 40% was accompanied

398

by an increase in the expression of CPSase and also by expression increases in the salvage

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enzymes UKase and UPRTase, these in turn, resulted in an increase in the pools of UTP, UDP,

400

and UDP-glucose up to 2-fold higher than wild type. The authors concluded that the wild type

401

uridine nucleotide pools limit the use of sucrose for starch and cell wall synthesis, and they

402

commented that it is surprising that mechanisms for decreasing de novo synthesis have not

403

developed during evolution or in the course of plant breeding, speculating that the pathway must

404

provide advantages under certain conditions (Geigenberger et al., 2005). In a contemporary

405

study on S. tuberosum and tobacco from Zrenner’s lab (Schröder et al., 2005), an antisense

406

strategy was used to produce potato plant lines with ATCase or DHOase proteins levels reduced

407

to < 50% of wild type. These plants showed decreased growth, but exhibited no pleiotropic

408

effects or alterations in the developmental program, and only slight changes in pyrimidine

409

nucleotide levels in mature tissues. Carbohydrate levels were decreased in roots, and increased

410

in leaves. Plants with < 10% of wild type DHOase protein eventually died without flowering or

411

tuberization. Tobacco plant lines with decreased UMP synthase expression were produced but

412

exhibited no phenotypic differences from wild type, however, the authors commented that the

413

reduction in the protein levels may not have been sufficient to cause such differences. They

414

concluded that the response of the plant to a moderate decrease in pyrimidine biosynthesis was to

415

decrease its growth rate in accordance with the size of the available nucleotide pool, however,

416

they noted that the size of the pool may depend on the type of tissue and its developmental stage

417

(Schröder et al., 2005). Support for this idea was provided by a later study, which demonstrated

418

dynamic regulation of the PYRB-encoded ATCase protein during Arabidopsis thaliana growth

419

and development as shown by monitoring the activity of a reporter gene under the control of the

420

PYRB promoter (Chen and Slocum, 2008). Similarly, Arabidopsis lines with RNAi-mediated 70

421

– 95% knockdowns of PYRB transcripts and concomitant reduced ATCase protein levels had

422

delayed growth and development compared with wild type plants (Chen and Slocum, 2008).

423

Plants having only 5% of the wild type expression produced seeds with extremely poor viability.

424

This Arabidopsis study is in general agreement with the results of Geigenberger and co-workers

425

(Geigenberger et al., 2005), for example, knockdown of a pyrimidine biosynthetic enzyme, in

426

this case ATCase, resulted in increased expression of another biosynthetic enzyme, in this case

427

UMP synthase, and increased expression of the salvage enzyme UPRTase (Chen and Slocum,

428

2008). The conclusions that can be drawn from these three studies are that plants dynamically

429

regulate synthetic and salvage pathways according to pyrimidine availability, and a decrease of

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430

flux through the former simply leads to a slower growth rate but does not result in phenotypic

431

abnormalities. However, the synthetic pathway is clearly essential, because decreasing activity

432

below certain thresholds results in loss of viability. Genetic manipulations of pyrimidine metabolism in multicellular organisms often have

434

complex effects that are not easy to dissect. Recent studies on UPase in mice illustrate this point

435

(Le et al., 2013; Urasaki et al., 2014). A knock-in of UPase resulted in enzyme overexpression

436

and a 25-fold increase in activity. The livers of the mice exhibited a 13-fold decrease in uridine

437

levels, and more than 2-fold higher levels of beta-alanine. The mice were found to suffer from

438

hepatic vesicular steatosis, which could be suppressed by administration of uridine or by

439

inhibition of dihydropyrimidine dehydrogenase. Changes in liver protein acetylation upon

440

administration of uridine were observed in mice and in primary hepatocyte cultures. Testing of

441

inhibitors and metabolites of de novo pyrimidine biosynthesis showed that brequinar, an inhibitor

442

of DHODase, and its product orotate, also caused hepatic vesicular steatosis (Le et al., 2013).

443

The authors attributed the effect of orotate to inhibition of DHODase and suggested that this

444

compromised mitochondrial respiration in the liver. Nevertheless, orotate is known to affect

445

expression levels of enzymes involved in lipid metabolism (Jung et al., 2011).

446

important to point out that the contribution of DHODase activity to electron transport flux or to

447

the function of respiratory complexes is still unknown. Furthermore, human genetic deficiencies

448

in UMP synthase, which lead to high levels of orotate causing orotic aciduria, and deficiencies in

449

DHODase (Miller syndrome), have very different clinical features (Balasubramaniam et al.,

450

2014). In another study, mice with a knockout of UPase (UPase-/-) were found to have uridine

451

increases of 6-fold in plasma and 3-fold in the liver (Le et al., 2013; Urasaki et al., 2014). They

452

exhibited a decrease in insulin-stimulated blood glucose removal, and an activation of the heme-

453

deficiency response pathway. Interestingly, although these last two effects were also observed in

454

wild type mice treated with short term supplementation of uridine, additional effects, for

455

example increased liver protein glycosylation and reduced phosphorylation of insulin signaling

456

proteins, were not observed in the knockout mice.

It is also

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457 458

(6) Concluding remarks.

459

In this review, we have presented several studies on pyrimidine metabolism in pathogen-

460

infected hosts. During their evolutionary development, pathogens have developed strategies for

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manipulating pyrimidine metabolism in the organisms they infect, thereby generating cellular

462

host environments that provide optimum levels of pyrimidine metabolites necessary for pathogen

463

proliferation. We would like to learn how these consummate manipulators of host cell

464

metabolism alter pyrimidine levels for their own ends. Perhaps such knowledge could be applied

465

to the ongoing challenge of understanding how pyrimidine inhibitors or antimetabolites can be

466

used to achieve a desired effect, for example to kill viruses, parasites, or other pathogens. Several

467

obstacles have impeded researchers from achieving these goals. A problem inherent in the study

468

of pathogen-infected hosts is that it is difficult or impossible in many cases to determine which

469

organism the metabolites originated from. Another problem is that information on the cellular

470

system under study is often incomplete, for example, expression data may be readily available,

471

but information on mRNA stability, metabolite levels, or enzyme activities may be lacking. The

472

integration and analysis of available data from new strategies, such as genomics, transcriptomics,

473

proteomics, metabolomics, is beginning to permit the construction of a more complete picture. It

474

should be noted that although these new techniques will permit a more comprehensive

475

description of the effect of an inhibitor or a genetic manipulation on the cell, the mechanisms

476

which led to that effect may still not be understood. Despite extensive biochemical knowledge

477

about pyrimidine metabolism, a global vision of its dynamics is only starting to emerge.

478 479

Acknowledgements

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This work was supported in part by the Universidad de los Andes Faculty of Sciences, by

481

basic sciences project P13.700022.007 from the University of los Andes Office of Research, by

482

COLCIENCIAS grant # 120-4521-28532. We would like to give a special acknowledgement to

483

Colciencias (Programa de Apoyo a Doctorados Nacionales) for the financial support of the Ph.D.

484

studies of M.F.G. and H.Y.N.-O. We regret that some important findings were not included here

485

due to the limited space.

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Sigoillot, F.D., Berkowski, J.A., Sigoillot, S.M., Kotsis, D.H., Guy, H.I., 2003. Cell cycledependent regulation of pyrimidine biosynthesis. J. Biol. Chem. 278, 3403–3409.

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Sreedharan, S., Shaik, J.H.A., Olszewski, P.K., Levine, A.S., Schiöth, H.B., Fredriksson, R., 2010. Glutamate, aspartate and nucleotide transporters in the SLC17 family form four main phylogenetic clusters: evolution and tissue expression. BMC Genomics 11, 17.

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Stasolla, C., Loukanina, N., Yeung, E.C., Thorpe, T. a, 2004. Alterations in pyrimidine nucleotide metabolism as an early signal during the execution of programmed cell death in tobacco BY-2 cells. J. Exp. Bot. 55, 2513–2522.

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Tiedje, K.E., Stevens, K., Barnes, S., Weaver, D.F., 2010. Beta-alanine as a small molecule neurotransmitter. Neurochem. Int. 57, 177–188.

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Urasaki, Y., Pizzorno, G., Le, T.T., 2014. Uridine affects liver protein glycosylation, insulin signaling, and heme biosynthesis. PLoS One 9, e99728.

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Vastag, L., Koyuncu, E., Grady, S.L., Shenk, T.E., Rabinowitz, J.D., 2011. Divergent effects of human cytomegalovirus and herpes simplex virus-1 on cellular metabolism. PLoS Pathog. 7, e1002124.

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Vogels, G and van der Drift, C., 1976. Degradation of purines and pyrimidines by microorganisms. Bacteriol. Rev. 40, 403–468.

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Webb, M.E., Smith, A.G., Abell, C., 2004. Biosynthesis of pantothenate. Nat. Prod. Rep. 21, 695–721.

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640

Fig. 1. Overview of human pyrimidine metabolic pathways.

642

Transporters important for pyrimidine metabolism are shown at the top of the figure. Human

643

cells can use transporters from the solute carrier family (SCL17) to exchange amino acids

644

(aspartate and glutamate) and nucleotides (Sreedharan et al., 2010), while nucleobases can exit or

645

enter the cell via the equilibrative nucleoside transporters (ENTs) and the concentrative

646

nucleoside transporters (CNT1) (Sahoo et al., 2014). Transporters of the ABC transport family

647

import nucleotides and nucleosides into the cell. The de novo pyrimidine biosynthetic pathway

648

includes six enzymatic steps (far left, purple). The first three steps are catalyzed by a

649

multifunctional protein called CAD and the last two steps by a bifunctional enzyme, UMP

650

synthase.

651

transferases (cyan). Nucleosides may be phosphorylated to form nucleoside monophosphates,

652

diphosphates and triphosphates by nucleoside kinases (pink and orange). Nucleoside

653

diphosphates can be transformed into deoxynucleosides by the ribonucleotide reductase system.

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Inside the cells, nucleobases are converted to nucleosides by phosphoribosyl

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The lower part of the figure shows that pyrimidine nucleotides are the main precursors for

655

synthesizing lipids (blue) and UDP-sugars (gray), and are also essential for the formation of

656

RNA (green) and DNA (brown). Pyrimidines are degraded to ß-alanine or ß-aminoisobutyrate

657

(far right, yellow). The latter can be further degraded to methylmalonate-semialdehyde which is

658

an intermediate in valine degradation. Enzymes are denoted by their Enzyme Commission (EC)

659

numbers: Dihydropyrimidine dehydrogenase (1.3.1.2); dihydroorotate dehydrogenase (1.3.5.2);

660

ribonucleoside-diphosphate reductase (1.17.4.1); thioredoxin-disulfide reductase (1.8.1.9);

661

thymidylate synthase (2.1.1.45); aspartate transcarbamoylase (2.1.3.2); uridine phosphorylase

662

(2.4.2.3); thymidine phosphorylase (2.4.2.4); uracil phosphoribosyltransferase (2.4.2.9); orotate

663

phosphoribosyltransferase (2.4.2.10); thymidine kinase (2.7.1.21); uridine kinase (2.7.1.48);

664

deoxycytidine kinase (2.7.1.74); nucleoside diphosphate kinase (2.7.4.6); thymidylate kinase

665

(2.7.4.9); adenylate kinase G (2.7.4.10); UMP/CMP kinase (2.7.4.14); RNA polymerase

666

(2.7.7.6); DNA polymerase (2.7.7.7); UDP-glucose pyrophosphorylase (2.7.7.9); ethanolamine-

667

phosphate cytidylyltransferase (2.7.7.14); choline-phosphate cytidylyltransferase (2.7.7.15);

668

UTP:N-acetyl-a-D-glucosamine-1-phosphate

669

cytidylyltransferase

670

pyrophosphatase (3.6.1.19); beta-ureidopropionase (3.5.1.6); dihydropyrimidinase (3.5.2.2);

671

dihydroorotase

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diphosphatase (3.6.1.6); dCTP pyrophosphatase 1 (3.6.1.12); deoxyuridine triphosphatase

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(3.6.1.23); orotidine 5-phosphate decarboxylase (4.1.1.23); cytidine triphosphate synthase

674

(6.3.4.2); carbamoyl phosphate synthase (6.3.5.5). Abbreviations: ADP, adenosine diphosphate;

675

Asp, aspartate; ATP, adenosine triphosphate; ß-Ala, ß-alanine; ß-AIB, ß-aminoisobutyrate; ß-

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UIB, ß-ureidoisobutyrate; ß-UP, ß-ureidopropionate; CDP, cytidine-5-diphosphate; CDP-Cho,

677

CDP-choline; CDP-Etn, CDP-ethanolamine; CMP, cytidine-5-monophosphate; CTP, cytidine-5-

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(2.7.7.41);

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(3.5.2.3);

cytidine

uridylyltransferase

5′-nucleotidase

deaminase

(3.1.3.5);

(3.5.4.5);

(2.7.7.23);

phosphatidate

nucleoside

triphosphate

apyrase

(3.6.1.5);

nucleoside

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triphosphate; dCDP, deoxy-cytidine-5-diphosphate; dCMP, deoxy-cytidine-5-monophosphate;

679

dCTP, deoxy-cytidine-5-triphosphate; DHF, dihydrofolate; DHT, dihydrothymine; DHU,

680

dihydrouracil;

681

monophosphate; dTTP, deoxy-thymidine-5-triphosphate; dUDP, deoxy-uridine-5-diphosphate;

682

dUMP, deoxy-uridine-5-monophosphate; dUTP, deoxy-uridine-5-triphosphate; Glu, glutamate;

683

Glu-1-P, glucose-1-phosphate; Gly, glycine; NADPH, nicotinamide adenine dinucleotide

684

phosphate; OMP, orotidine-5-monophosphate; Pi, inorganic phosphate; PPi, pyrophosphate;

685

PRPP, phosphoribosyl pyrophosphate; Ser, serine; UDP, uridine-5-diphosphate; UMP, uridine-5-

686

monophosphate; UQ, ubiquinone; UTP, uridine-5-triphosphate; THF, tetrahydrofolate.

687

diagram is based on data from the Kyoto Encyclopedia of Genes and Genomes (KEGG)

688

Pathway, hsa00240 (http://www.genome.jp/kegg-bin/show_pathway?map00240). ChemAxon

689

software was employed to build the chemical structures.

690 691

Fig. 2. A heat map illustrating changes in the expression of the genes of pyrimidine metabolism

692

in pathogenic eukaryotes.

693

A graphical heat map representation was constructed to illustrate changes (log2) in expression

694

among two contrasting time points or conditions important in the infection process of selected

695

plant and animal pathogens. Only RNA Sequencing (RNA-seq) data were used for its

696

construction since these represent the quantity of the transcripts at a given moment or condition.

697

Red and green color intensities indicate increases and decreases, respectively, in the gene

698

expression profile. Gray indicates the absence of the gene. Toxoplasma gondii: data were

699

obtained for tachyzoites of the RH strain from infected human foreskin fibroblasts from ToxoDB

700

13.0. Time points were early in infection, 2 hpi, and late in infection, 20 hpi. Plasmodium

701

falciparum: data were obtained for strain 3D7 during intra-erythrocytic development from

702

PlasmoDB 9.0. Selected time points were 20 hpi (trophozoite) and 40 hpi (schizont).

703

Cryptosporidium parvum: data were obtained for sporozoites of isolate IPZ:CH-Crypto_K6769

704

infection of CT-8 cells from CryptoDB 7.0. The time points selected were infected cells

705

incubated for 48 hpi and 96 hpi. Trypanosoma brucei: data were obtained for strain Lister 427

deoxy-thymidine-5-diphosphate;

dTMP,

deoxy-thymidine-5-

The

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dTDP,

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from TriTrypDB 8.1. The comparison was generated between proliferative bloodstream stage in

707

the mammalian host, and the procyclic form, from which it differentiates, in the midgut of the

708

tsetse fly. Cryptococcus neoformans: data were obtained for variety grubii H99 from wild-type

709

data from the FungiDB 3.1. The comparison was generated between capsule inducing (37°C with

710

CO2) and non-inducing (30°C) conditions. Colletotricum higginsianum: a pathogenic plant

711

fungus, data were obtained for the in planta stages during infection of Arabidopsis thaliana leaf

712

sheaths from the Gene Expression Omnibus GSE33683. Selected time points were 22 hpi,

713

representing the earliest stage of biotrophy with penetrating apressoria, and 60 hpi in the

714

necrotrophic phase. Phytophthora ramorum: data were obtained from the FungiDB 3.1. The

715

comparison was generated between (sporangia+zoospores) germination stages and non-

716

germinating mycelia grown in tomato broth. In addition, we present expression data for an

717

infected host, Mus musculus (mouse macrophages): data were obtained for primary bone

718

marrow-derived macrophages from C57BL/6 mice infected T. gondii RH strain from HostDB

719

1.1. Time points were uninfected macrophages and infected macrophages 22 hpi. Viewing was

720

done with Mev 4.8.1.

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Supplementary Table 1. Analyses of the genes of pyrimidine metabolism in selected eukaryotic genomes.

✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

✓ ✓ ✓ ✓ ✓ ✓ ✓

✓ ✓ ✓ ✓ ✓ ✓ ✓

✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

Low mRNA expression levels Exclusively for IMP/GMP Replaced by 3.6.1.6 Nucleoside diphosphate phosphatase (no information about whether it is purine or pyrimidine specific) Gene exists but is inactive Absent

✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

✓ ✓ ✓ ✓ ✓ ✓

✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

2.4.2.3 Uridine phosphorylase

3.5.4.5 Cytidine deaminase

3.1.3.5 5'-nucleotidase

3.6.1.5 Apyrase

3.6.1.12 dCTP pyrophosphatase

2.7.1.21 Thymidine kinase

2.7.4.9 Thymidylate kinase

2.7.1.74 Deoxycytidine kinase

2.7.1.48 Uridine kinase

2.7.4.6 Nucleoside diphosphate kinase

2.7.4.14 UMP/CMP kinase

✓ ✓ ✓ ✓ ✓

3.6.1.19 Nucleoside triphosphate pyrophosphatase

✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

RI PT

✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

SC

2.1.1.45 Thymidylate synthase ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

2.4.2.10 Orotate phosphoribosyltransferase

1.3.5.2 Dihydroorotate dehydrogenase

6.3.4.2 Cytidine triphosphate synthase

✓ ✓ ✓

✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓ ✓

TE D

3.5.2.3 Dihydroorotase ✓ ✓ ✓ ✓ ✓ ✓ ✓

EP

✓ ✓ ✓ ✓

4.1.1.23 Orotidine 5-phosphate decarboxylase

✓ ✓ ✓

✓ ✓ ✓ ✓

2.1.3.2 Aspartate transcarbamoylase

6.3.5.5 Carbamoyl phosphate synthase

3.5.1.6 Beta-ureidopropionase

✓ ✓ ✓ ✓

AC C

Cryptosporidium parvum Plasmodium falciparum Toxoplasma gondii Trypanosoma brucei Phytophthora ramorum Candida albicans Ustilago maydis Cryptococcus neoformans Caenorhabditis elegans Arabidopsis thaliana Homo sapiens

3.5.2.2 Dihydropyrimidinase

Organism

1.3.1.2 Dihydropyrimidine dehydrogenase

E.C number - Enzyme

2.4.2.9 Uracil phosphoribosyltransferase

Salvage & Modifications 1.17.4.1 Ribonucleoside-diphosphate reductase

Synthesis

M AN U

Catabolism

✓ ✓ ✓ ✓ ✓

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Supplementary Table 1. Analyses of the genes of pyrimidine metabolism in selected eukaryotic genomes.

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The table highlights the differences in the genomic metabolic repertoire of the pyrimidine pathways among the eukaryotes mentioned in this review. Presence or absence of the sequences of the enzymes of pyrimidine metabolism was evaluated for each organism using KEGG, accessing by gene code, name, and EC number. A second validation was performed with the same criteria but using instead the data available in each organism’s specific genome database. For genes not present, possible misannotation problems were further evaluated by analyzing the genomes to look for hypothetical proteins that shared the conserved protein domain of the target protein. A check mark represents the presence of coding sequence for a protein. Enzymes are grouped as pathways of catabolism, synthesis, salvage and modification. Enzyme numbers are the same as in Figure 1. Specific genomic data were retrieved from: Cryptosporidium parvum: http://cryptodb.org; Plasmodium falciparum: http://plasmodb.org; Toxoplasma gondii: http://toxodb.org; Trypanosoma brucei: http://tritrypdb.org; Phytophthora ramorum: http://genome.jgi-psf.org/; Candida albicans: http://www.candidagenome.org; Ustilago maydis: http://www.broadinstitute.org; Cryptococcus neoformans: http://www.broadinstitute.org; Caenorhabditis elegans: http://www.wormbase.org; Arabidopsis thaliana: http://www.arabidopsis.org/index.jsp; Homo sapiens: http://www.genome.jp/kegg-bin/show_pathway?org_name=hsa&mapno=00240&mapscale=&show_description=show.