Carbon Chapter 125 UTP

Virology

Volume 40, Issue 2, February 1970, Pages 244-250

Properties of the products of UTP incorporation by cell-free extracts of leaves infected with bromegrass mosaic virus or with broadbean mottle virus

Author links open overlay panelJ.Semal

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Abstract

After deproteinization with phenol and detergent, the product of UTP-3H incorporation by a particulate fraction prepared from barley leaves infected with bromegrass mosaic virus displayed properties consistent with those of a double-stranded RNA. The major part of the label incorporated in the product was resistant to pancreatic ribonuclease (RNase) (5 μg/ml) in 1 × SSC, but was made acid-soluble by treatment with RNase in 0.1 × SSC. Similar results were obtained with the deproteinized product of UTP-3H incorporation by extracts from broadbean leaves infected with broadbean mottle virus. When the deproteinized product synthesized by extracts from bromegrass mosaic virus-infected leaves was heated in 1 × SSC at increasing temperatures, a sharp transition from RNase resistance to RNase sensitivity was observed, with a Tm of 99–100°.

With both bromegrass mosaic virus and broadbean mottle virus, the particle-bound product of UTP-3H incorporation was double-stranded in its native state, as suggested by RNase resistance in 1 × SSC and RNase sensitivity in 0.05 × SSC, respectively, of the radioactive product associated with the particulate leaf fraction. The native products of UTP-3H incorporation were sensitive to RNase in 0.05 × SSC without any pretreatment of the leaf extracts.

Abstract

Increasing evidenced for receptors for uracil nucleotides has focused interest on specific signalling mechanisms involving UTP and UDP. At least three metabotropic P2 receptors are stimulated by uracil nucleotides with equal or greater potency than by adenine nucleotides, and there might be ionotropic receptors as well. Regulation of uridine and uracil nucleotide levels is important when considering the receptor-mediated effects of these compounds. Cells can synthesize uracil nucleotides de novo or by salvage of uridine. UTP made from salvage might be preferentially unsed for RNA synthesis in the nucleus, while UTP synthesized de novo seems to be used for UDP-sugar and CDP-phospholipid production. UTP from both pathways can enter a free UTP pool, from which UTP can be released from cells. UTP and UDP can stimulate pyrimidinoceptors, but metabolism by ectonucleotides limits their effects. Alternatively, UTP might be a substrate for ecto-protein kinases,and these could contribute to its extracellular regulation. Cells can reclaim uridine, using nucleoside transport processes, following dephosphorylation of UTP, UDP and UMP. In this article Christopher Anderson and Fiona Parkinson discuss how understanding the processes that regulate uridine and uracil nucleotide concentrations with enhance our ability to manipulate UTP/UDP signalling pathways for pharmacological effect.

1 Release of UTP

Cellular release of UTP was first reported in studies indicating that perfused [3H]uridine-labeled endothelial cells released [3H] species in response to changes in flow rates (Saïag et al., 1995). Taking advantage of the high selectivity that UDP-glucose pyrophosphorylase exhibits for the conversion of UTP to UDP-glucose in the presence of glucose-1 phosphate, an assay that quantifies the UTP-dependent conversion of [14C]glucose-1P to [14C]UDP-glucose with nanomolar sensitivity was developed (Lazarowski & Harden, 1999). Using this assay, the mass of UTP released from various cell types and tissues was quantified under various physiological conditions. It was thus established that, like ATP, UTP is released from cells constitutively and that stimuli promoting ATP release also result in enhanced release of UTP. In most (but not all) cases where UTP release has been measured, the extracellular UTP:ATP mass accumulation ratio (~ 1:3) resembles the intracellular UTP:ATP ratio, suggesting that both nucleotides were released via common mechanisms and from a common subcellular nucleotide pool (Lazarowski & Harden, 1999). Enhanced release of UTP has been observed with various cultured cell types subjected to mechanical stresses (Lazarowski & Harden, 1999; Lazarowski et al., 1997; Tatur et al., 2008), as well as in apoptotic T lymphocytes (Elliott et al., 2009), lung secretions from RSV-infected mice (Davis et al., 2006), and the blood following heart ischemia (Erlinge et al., 2005). Davis et al. illustrated that removal of UTP from the bronchoalveolar liquid of RSV-infected mice in vivo markedly improved alveolar fluid clearance (Davis et al., 2004, 2006). These observations suggest that UTP secreted into the alveolar space plays a role in the pathogenesis of lung edema associated with viral infection. The recent observation that UTP was more efficient than ATP in promoting Cl− secretion from primary cultures of alveolar type II cells further supports the involvement of UTP release in alveolar epithelial cell homeostasis (Bove et al., 2010).

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G Protein Coupled Receptors

Kenneth A. Jacobson, ... Francesca Deflorian, in Methods in Enzymology, 2013

3 Receptors for Extracellular Nucleotides and their Modeled Structures

ATP, UTP, and other nucleotides are released from intracellular sources in response to stress (such as hypoxia, ischemia, or mechanical stress) as a result of tissue damage, through vesicular release as a cotransmitter or through pannexin hemichannels. Upon release, these nucleotides can activate P2X channels or P2YRs, which tend to mobilize a response to the challenge, such as intensifying the immune response. Over time, through the action of nucleotidases on adenine nucleotides, adenosine is produced, or it may also originate from cellular release.

The P2YRs belong to a branch of class A GPCRs chiefly composed of receptors for nucleotides, lipids, metabolites of the Krebs cycle, and protease-activated receptors (Costanzi et al., 2004). Second messenger coupling and phylogenetic analyses based on sequence comparisons revealed that the P2YRs can be subdivided into two distinct groups: the P2Y1-like subfamily, comprising the P2Y1, P2Y2, P2Y4, P2Y6, and P2Y11 subtypes, and the P2Y12-like subfamily, comprising the P2Y12, P2Y13, and P2Y14 subtypes (Table 9.1). The missing numbers are due to earlier erroneous attribution of certain receptors, which in fact are not activated by nucleotides, to the P2Y family. The sequence identity is significantly high between members of the same subfamily, while it is relatively low across the two subfamilies (from 14% for P2Y1R/P2Y14R to 50% for P2Y2R/P2Y4R). Notably, the classification based on sequence alignment coincides with a pharmacological division based on the coupling of the receptors to different G proteins: the members of P2Y1-like subfamily activate the phospholipase C (PLC) signaling pathway via Gq, while those of the P2Y12-like subfamily inhibit the adenylyl cyclase pathway via Gi.

Experimentally elucidated structures are not yet available for any of the P2YRs. In the absence of such structures, in the past decade, we have constructed models of the members of this family of receptors based on the various available templates. The earlier models were all based on rhodopsin, which, at the time, was the only crystallographically solved GPCR (Ivanov et al., 2006). We incorporated into the model experimentally derived hypotheses on the structures of the P2YRs. These included the presence of a disulfide bridge that putatively connects the third transmembrane domain (TM3) with the second EL in virtually all GPCRs and a second disulfide bridge between the N-terminal domain with EL3 that was proposed, on the basis of mutagenesis studies conducted at the P2Y1 and P2Y2Rs, as a constraint to help characterize the extracellular region. Our models also incorporated a salt bridge connecting EL2 of P2Y1R with the extracellular end of TM6, which was supported by experimental data. Specifically, data from mutagenesis studies revealed that the residues involved in formation of these extracellular bridges are fundamental to receptor function.

On the basis of these rhodopsin-based in silico structures, through molecular docking experiments, we generated models of the complexes of the P2YRs with both natural and synthetic nucleotides ligands, including agonists and antagonists (Table 9.1). Experimentally derived data were amply used to guide the molecular docking. Most notably, modeling-guided mutagenesis data revealed two distinct sets of cationic residues in the P2Y1R- and the P2Y12R-like subfamilies to be implicated in the recognition of the phosphate moieties of the nucleotides. Three cationic residues were identified at positions 3.29 of TM3, 6.55 of TM6, and 7.39 of TM7 for the members of the P2Y1R-like subfamily and at positions EL2.52 of EL2, 6.55 of TM6, and 7.43 of TM7 for the P2Y12R-like subfamily. Our molecular models of the P2YRs in complex with their ligands were constructed by anchoring the phosphates of the nucleotides to these key cationic residues (Costanzi et al., 2004).

The puckering conformation of the sugar moiety of the ligands adopted in our models was also experimentally supported. Specifically, the ribose conformation necessary for the recognition of nucleotides by each of the P2Y1R-like subtypes was inferred through the synthesis of rigid methanocarba analogues of the ribonucleotides, constrained in two isomeric forms as either the northern (N) or the southern (S) conformation by the fusion of a cyclopentane and a cyclopropane ring (Costanzi et al., 2005), and most of these subtypes preferred to bind with their ligands in the (N) conformation. The only exception was the P2Y6R, which recognized the sugar moiety of its ligands only in the (S) conformation. Notably, this eccentric characteristic of the P2Y6R was strikingly anticipated by our models based on Monte Carlo searches followed by molecular dynamics simulations.

More recently, the structure of the CXCR4 chemokine receptor was solved crystallographically (Wu et al., 2010). Among the currently crystallized GPCRs, this receptor is the most suitable template for the modeling of the P2YRs. As we previously illustrated, CXCR4 is a member of a branch of peptide-activated GPCRs that appears to be phylogenetically the closest to the P2Y branch (Deflorian & Jacobson, 2011). Among the crystallographically solved GPCRs, this receptor shares the highest sequence similarity with P2YRs (about 25% calculated relatively to the entire portion of the CXCR4 receptor that was solved crystallographically, with the exclusion of the C-terminal domain) (Maruoka et al., 2011). Moreover, the CXCR4 receptor contains the above-mentioned second disulfide bridge connecting the N-terminus (through a Cys located two positions upstream when compared to the P2Ys) with a Cys situated between EL3 and TM7 (at position 7.25, as in the P2YRs). Before the CXCR4 structure was solved, none of the structurally characterized GPCRs featured a similar disulfide bridge.

For these reasons, we built new models of the members of the P2Y family based on this new peptide receptor template. Different conformations of the EL2 domain are immediately apparent when comparing the models of a P2YR based on either rhodopsin or CXCR4 (Fig. 9.3). Specifically, the rhodopsin-based models feature a loop that occludes like a plug the extracellular opening of the interhelical binding cavity. On the contrary, the CXCR4-based models feature a more solvent-exposed EL2 domain that leaves the interhelical binding cavity more open toward the extracellular milieu. Moreover, the choice of receptor template for modeling of the P2Y4R had major effect on the position of the pharmacophore, in either of two ligands: UTP or a selective agonist, MRS4062. Such a difference is probably attributable to the peculiarity of the biology of rhodopsin when compared to the CXCR4 receptor and the great majority of GPCRs. Unlike most GPCRs, whose ligands diffuse from the extracellular space into the receptor, rhodopsin features a ligand already covalently bound within the interhelical cavity.

Figure 9.3. Side-by-side view of two P2Y4R models (Maruoka et al., 2011): an older rhodopsin-based model is shown in (A), while a more recent CXCR4-based model is shown in (B). The two models can be readily recognized by the distinctive structure of their second extracellular loop (EL2) domains—the beta-hairpins represented with thick gray antiparallel arrows. Specifically, in the rhodopsin-based model EL2 lays low over the extracellular opening of the helical bundle, while in the CXCR4-based model it projects toward the extracellular space. As a result, the docked ligand is pushed more deeply toward the center of the receptor in the first than in the latter. The structure of the receptors is schematically represented as a ribbon with a color gradient going from blue at the N-terminus to red at the C terminus (TM1: dark blue; TM2: cyan; TM3: green; TM4: yellow/green; TM5: yellow; TM6: orange, TM7: red). The ligands—UTP in the rhodopsin-based model and an N4-substituted derivative of CTP (MRS4062, N4-(3-phenylpropoxy)-cytidine-5′-triphosphate) in the CXCR4-based model—are represented as van der Waals spheres (orange, P; red, O).

Notably, the new CXCR4 receptor-based P2YR models explain ligand SAR more effectively than those based on rhodopsin, suggesting that they may be closer to the actual structure of the receptors. For instance, our new models of the pyrimidine-nucleotide binding P2Y2 and P2Y4Rs, unlike their rhodopsin-based counterparts, were in agreement with the activity and selectivity profile of compounds bearing relatively large substituents attached to the nucleobase (Maruoka et al., 2011). Moreover, a model of the P2Y12R also based on the CXCR4 receptor was revealed to be in excellent agreement with the SAR of both agonists and antagonists (Deflorian & Jacobson, 2011).

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Pharmacology of Purine and Pyrimidine Receptors

Ivar von Kügelgen, T. Kendall Harden, in Advances in Pharmacology, 2011

B P2Y2 Receptor

Triphosphate nucleotides including UTP, ATP, UTPγS, and ATPγS act as full agonists at this receptor (Table I). 2-Thio-UTP and its 2′-amino-2′-deoxy-analog (MRS2698) are selective P2Y2-receptor agonists (El-Tayeb et al., 2006; Ivanov et al., 2007a; Jacobson et al., 2006). In addition to triphosphate nucleotides, the receptor also responds to dinucleoside polyphosphates including diadenosine-tetraphosphate (Ap4A; Lazarowski et al., 1995; Patel et al., 2001) as well as to Up4U (diquafosol, INS365; Pendergast et al., 2001). INS365 is undergoing phase III clinical trials for the treatment of dry eye disease (Tauber et al., 2004). The dinucleoside analog P1-(uridine 5′)-P4-(2′-deoxycytidine-5′) tetraphosphate (INS37217) is a potent agonist at the P2Y2 receptor with some effects at the P2Y4 receptor (Table I).

Suramin blocks the P2Y2 receptor with an affinity about 20 times lower than observed at the P2Y1 receptor (Table II). Reactive blue-2 acted as an antagonist at the recombinant human P2Y2 receptor with an IC50 of 2 μM (Hillmann et al., 2009). Flavonoid derivatives and reactive blue-2 derivatives have been studied for their antagonistic action at the P2Y2 receptor (Brunschweiger & Müller, 2006; Weyler et al., 2008). Among these compounds, the reactive blue-2 analog PSB416 displayed an IC50 of 22 μM (Hillmann et al., 2009). Some acyclic nucleotide analogs of UTP showed antagonistic properties at the P2Y2-receptor; however, their affinities are low (Sauer et al., 2009).

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Caenorhabditis elegans: Cell Biology and Physiology

Lois G. Edgar, Bob Goldstein, in Methods in Cell Biology, 2012

V Radioactive Labeling

Both UMP and UTP labeled with 3H or 32P are readily taken up by embryos after removal of the vitelline envelope, and can be visualized by autoradiography (Edgar et al., 1994); one supposition as to how the triphosphate gets in is that cell membranes may be slightly leaky.

Radioactive labeling requires extreme care, as working with small numbers of embryos must be done under the dissecting scope and it is virtually impossible to avoid all mouth pipetting. Likewise, wearing gloves makes the manipulation very difficult and probably increases the chance of contamination. It can be done most safely using the Cel-Line slides and moving embryos in small batches from drop to drop. A plastic shield should be fitted around the dissecting microscope for labeling with 32P. Appropriate safety precautions should be taken. The mouth pipette should be fitted with a cotton plug or a very long tube, and care should be taken never to overfill the glass tip.

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Pharmacology of Purine and Pyrimidine Receptors

Filip Kukulski, ... Jean Sévigny, in Advances in Pharmacology, 2011

2 Effect on Vasoconstriction

Either extracellular UDP or UTP induces a contractile response via P2Y6 receptor in mouse aortic rings denuded from its endothelium. In agreement with an essential role of NTPDase1 in the control of P2Y6-mediated vasoconstriction, the weak response in wild-type animals was markedly enhanced in aortic rings from Entpd1−/− animals treated with UDP or UTP (Kauffenstein et al., 2010a). Accordingly, UDP infusion in vivo increased blood pressure more importantly in NTPDase1-deficient mice. In addition, Entpd1−/− mesenteric arteries displayed an enhanced myogenic response. This suggests that, in the absence of hydrolysis by NTPDase1, endogenous nucleotides released after mechanical forces, such as stretching in this case, can induce a more potent contraction in Entpd1−/− vessels.

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Ribonucleases - Part B

Agamemnon J. Carpousis, ... Vanessa Khemici, in Methods in Enzymology, 2001

Procedure

[32P]UTP-labeled 9Sa RNA is synthesized by transcription with T7 RNA polymerase of a DNA template prepared by PCR amplification.2 A detailed procedure is beyond the scope of this chapter. Sambrook et al.13 should be consulted for a general description of these procedures.

The following steps are performed on ice unless otherwise noted. Enough substrate for all the reactions is prepared by diluting [32P]UTP-labeled 9Sa RNA into SB (0.5–1.0 µ Ci/ml, final). Fractions from the large-scale purification are diluted 5-fold (lysate, S-30, S-200), 10-fold (ASP-40), or 20-fold (S-Sph, G-Grd) into EB. The enzyme is then further diluted in a series of 2-fold steps by mixing equal volumes of enzyme and EB, and changing the pipette tip after each dilution. Assemble the reactions on ice by mixing 2µl of enzyme with 8 µl of substrate and then incubating at 30° for 30 min. Quench the reactions on ice, add 10 µl of PK, and incubate at 50° for 10 min. Cool the reactions to room temperature, add 20 µl of FUD, heat at 85° for 5 min, and then quench on ice. A sample (20 µl) of each reaction is separated by electrophoresis on denaturing polyacrylamide gels. The gels are dried and exposed on X-ray film at −70° with an intensifying screen.

The autoradiograph in Fig. 2 shows the analysis of the S-200, ASP-40, and SP-Sph fractions from a large-scale purification. By careful visual inspection, it is possible to estimate 1 unit of activity. For example, in lanes 14–19 (SP-Sph fraction), partial digestion was observed in lanes 17–19 whereas in lane 16 the 9Sa substrate is completely cleaved at the a site and some of the 5′-a product is further digested by cleavage at the c site. Taking the amount of protein assayed in lane 16 (16 ng) as 1 unit of activity gives a specific activity of 62.5 units/µg of protein (62,500 units/mg). It is admittedly more difficult to make this determination with a crude fraction (S-200) because there is some undigested 9Sa substrate even at the highest concentration of protein. This leads to a systematic underestimation of RNase E activity that is apparently due to inhibition in the crude fractions. Whether the inhibitor(s) interferes directly with RNase E or acts indirectly, perhaps by sequestering the 9Sa substrate, is not known. An additional complication in assaying the crude fractions is that the yield of products does not appear to be quantitative. This could be due to degradation by other nucleases.

Fig. 2. Autoradiogram of a denaturing polyacrylamide gel [9% (w/v), 37.5 : 1,7 M urea, l Χ TBE showing the assay of RNase E activity in the S-200, ASP-40, and SP-Sph fractions from a large-scale purification of the degradosome (see Table I). The fractions were 2-fold serially diluted as described in text. The amount of total protein in the assay ranged from 6.4 to 0.20 µg (lanes 2 to 7), from 1.6 to 0.05 µg (lanes 8 to 13), and from 64 to 2 ng (lanes 14 to 19). Lane 1 is a control in which the 9Sa substrate was incubated under the same conditions and processed with the other samples. To the right is shown the position of the 9Sa substrate and the 5′-a, a-3′, 5′-c, and c-a products.

Table I shows an analysis of the amount of protein, activity, and specific activity in each step of a large-scale purification. Note that the activity increases between the lysate and ASP-40 fraction. This effect, which has been reproduced many times, is likely due to inhibition of RNase E activity in the crude fractions (discussed above). Because the activity is underestimated, we cannot calculate the yield and extent of purification in the initial fractions. We estimate that the yield of RNase E in the ASP-40 fraction is 40% on the basis of Western blot analysis. This gives an 8-fold enrichment in the ASP-40 fraction. The overall purification summarized inTable I shows a 3% yield with a 310-fold enrichment. These values seem reasonable given that the abundance of RNase E has been estimated as 1000 molecules per cell.15

Table I. RNase E Activity during Purification of Escherichia coli RNA Degradosomea

FractionProtein(mg)Activity(units)Specific activityYieldb(%)Enrichment (fold)Lysate18,000620,00034——S-309,800470,00048——S-2006,2001,600,000260——ASP-409002,300,0002,600408SP-Sph9.2670,00073,00012230G-Grd1.8180,000100,0003310

aPreparation starts with 100 g of cells. The amount of protein was determined by the method of Lowry, using a BSA standard. The activity was determined as described in text. The specific activity is expressed as units per milligram of protein. The fractions are lysate, total lysate before centrifugation; S-30, 30,000 g supernatant; S-200, 200,000 g supernatant; ASP-40, ammonium sulfate pellet (40% saturation); SP-Sph; chromatography on SP-Sepharose; and G-Grd, sedimentation on a glycerol gradient. Data presented are a composite of two preparations because we rarely assayed the lysate and S-30 fractions during the later stages of developing this procedure.bThe yield in the S-30 and S-200 fractions cannot be determined because the activity is inhibited in the crude fractions (see text). The 40% yield in the ASP-40 fraction is an estimation based on Western blotting.

Figure 3 shows an SDS–PAGE analysis of the fractions from a large-scale purification. To the right is shown the position of RNase E (Rne) and the three other major components of the RNA degradosome: Pnp (PNPase), Rh1B, and Eno (enolase). Rne* is a proteolysis product in which part of the C-terminal region of the protein has been removed. RNase E is known to be sensitive to proteolysis.2,16 Despite extensive precautions to minimize proteolysis, we have not been able to eliminate the Rne* product from the highly purified preparations of the RNA degradosome. Nevertheless, we believe that this proteolysis occurs during the purification because Rne* is not detected by Western blotting when total lysates are prepared by directly breaking the cells with heat in SDS–PAGE loading buffer.

Fig. 3. An SDS-polyacrylamide gel (9%, w/v) stained with Coomassie blue showing protein from various fractions of the large-scale purification. The amount of prote0in loaded in lanes 1 to 5 was, respectively, 74, 56, 32, 9.2, and 4.8 µg. To the right is indicated the position of RNase E (Rne), a proteolysis product of RNase E (Rne*), PNPase (Pnp), Rh1B, and enolase (Eno). Also indicated is a region of the gel where trace amounts of DnaK and polyphosphate kinase (Ppk) have been identified. RNase E is a 118-kDa protein that migrates at 180 kDa in SDS–PAGE because of the unusual amino acid composition of the C-terminal half of the protein. The molecular masses of the other major components by SDS–PAGE are PNPase (85 kDa), Rh1B (50 kDa), and enolase (48 kDa).

Just below Pnp there are several proteins in the 60- to 70-kDa range that are present in trace amounts in the G-Grd fraction. Among these are DnaK and polyphosphate kinase (Ppk), which have been identified by protein sequencing and for which there is evidence suggesting a physical association with the RNA degradosome.9,17

In Fig. 3 RNase E, which migrates as one of the largest proteins in E. coli, is clearly visible in the ASP-40 fraction. The only other protein identified in this region of the gel is MukB, which is 6-fold less abundant than RNase E.15 We have occasionally monitored the purification of the RNA degradosome, particularly during the later steps in the procedure, following RNase E by SDS–PAGE.

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Nucleotide Metabolism

N.V. Bhagavan, Chung-Eun Ha, in Essentials of Medical Biochemistry (Second Edition), 2015

Formation of Other Pyrimidine Nucleotides

UMP is the parent compound in the synthesis of cytidine and deoxycytidine phosphates and thymidine nucleotides (which are deoxyribonucleotides).

Synthesis of Cytidine Nucleotides

CTP is synthesized from UTP by transfer of the amide nitrogen of glutamine to C-4 of the pyrimidine ring of UTP. This reaction requires ATP as an energy source. See the following reaction:

Deoxycytidine phosphates result from reduction of CDP to dCDP by a mechanism analogous to that described for purine nucleotides. Then dCDP is converted to dCTP by nucleoside diphosphate kinase.

Synthesis of Thymidine Nucleotides

De novo synthesis of thymidylic acid (TMP) occurs exclusively by methylation of the C-5 of dUMP (Figure 25.18) by thymidylate synthase. The methylene group of N5,N10-methylene FH4 is the source of the methyl group, and FH4 is oxidized to FH2. For sustained synthesis, FH4 must be regenerated by dihydrafolate reductase. Recall that deoxynucleotides are formed at the diphosphate level by ribonucleotide reductase: thus, UDP is converted to dUDP; then to dUTP; dUMP is then generated mainly by dUTPase.

Figure 25.18. Synthesis of thymidylic acid (TMP). Fluorodeoxyuridylate inhibits conversion of dUMP to TMP, and methotrexate inhibits regeneration of the tetrahydrofolate coenzyme.

dUTP→dUTPasedUMP+PPi

The dUTPase reaction is very important because the DNA polymerases cannot distinguish dUTP from TTP and catalyze significant incorporation of dUMP into DNA when dUTP is present. Incorporation of the base U into DNA is not deleterious because all cells contain a uracil N-glycosylase that removes U from DNA. Figure 25.19 summarizes the pathway for TTP synthesis.

Figure 25.19. Pathway for TTP synthesis. The key intermediate is dUTP, which is converted to dUMP by dUTPase.

Thymidylate synthase is competitively inhibited by fluorodeoxyuridylate (FUDRP), with formation of a stable ternary complex with methylene FH4. FUDRP is generated by a salvage pathway from exogenous 5-fluorouracil (FU) or fluorodeoxyuridine (FUDR). FUDR is a useful drug in cancer chemotherapy because it inhibits TMP formation in proliferating cells. Thymidine nucleotide deficiency can also be induced by competitive inhibitors of dihydrofolate reductase, e.g., aminopterin (4-aminofolate) and methotrexate (4-amino-10-methylfolate; see Figure 25.3). 5-FU is normally inactivated by dihydropyrimidine dehydrogenase (DPD), up to 85% of the administered drug. Patients with inherited deficiency of DPD treated with standard doses of 5-FU experience serious clinical consequences leading to myelosuppression and gastrointestinal and neurologic toxicity. A similar toxic manifestation due to deficiency of thiopurine metabolizing enzyme was discussed earlier. The study of the role of genetic inheritance that leads to variations in drug response is known as pharmacogenomics.

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Nucleotide Metabolism

N.V. BHAGAVAN, in Medical Biochemistry (Fourth Edition), 2002

Formation of Other Pyrimidine Nucleotides

UMP is the parent compound in the synthesis of cytidine and deoxycytidine phosphates and thymidine nucleotides (which are deoxyribonucleotides).

Synthesis of Cytidine Nucleotides

CTP is synthesized from UTP by transfer of the amide nitrogen of glutamine to C-4 of the pyrimidine ring of UTP. This reaction requires ATP as an energy source.

The deoxycytidine phosphates result from reduction of CDP to dCDP by a mechanism analogous to that described for the purine nucleotides. Then dCDP is converted to dCTP by nucleoside diphosphate kinase.

Synthesis of Thymidine Nucleotides

De novo synthesis of thymidilic acid (TMP) occurs exclusively by methylation of the C-5 of dUMP (Figure 27-28) by thymidylate synthase. The methylene group of N5, N10-methylene FH4 is the source of the methyl group, and FH4 is oxidized to FH2. For sustained synthesis, FH4 must be regenerated by dihydrofolate reductase. Recall that deoxynucleotides are formed at the diphosphate level by ribonucleotide reductase; thus, UDP is converted to dUDP, then to dUTP, dUMP is then generated mainly by dUTPase.

FIGURE 27-28. Synthesis of thymidylic acid (TMP). Fluorodeoxyuridylate inhibits conversion of dUMP to TMP, and methotrexate inhibits regeneration of the tetrahydrofolate coenzyme.

dUTP→dUTPasedUMP+PPi

FIGURE 27-29. Pathway for TTP synthesis. The key intermediate is dUTP, which is converted to dUMP by dUTPase.

Because there is only one pathway for synthesis of TMP, it can be used to synthesize radioactively labeled DNA or to inhibit DNA synthesis selectively. In bacterial thymidylate synthase mutants (thy–), DNA synthesis is not possible without added thymine or thymidine, both of which can be utilized by salvage pathways. Neither thymine nor thymidine engages in synthetic reactions other than production of TMP, so radioactive thymine or thymidine can serve as unique precursors for synthesis of radioactive DNA. The radioactivity usually is in the methyl group; since this group is not metabolized, the appearance of radioactivity in any other compound is avoided.

Thymidylate synthase is competitively inhibited by fluorodeoxyuridylate (FUDRP), with formation of a stable ternary complex with methylene FH4. FUDRP is generated by salvage from exogenous 5-ftuorouracil (FU) or fluorodeoxyuridine (FUDR). FUDR is a useful drug in cancer chemotherapy because it inhibits TMP formation in proliferating cells. Thymidine nucleotide deficiency can also be induced by competitive inhibitors of dihydrofolate reductase, e.g., aminopterin (4-aminofolate) and methotrexate (4-amino-10-methylfolate; see Figure 27-3).

Fluoropyrimidines are catabolized by enzymes that normally participate in the breakdown of endogeneous pyrimidines uracil and thymine (see page 643). The deficiency of the initial rate-limiting enzyme dihydropyrimidine dehydrogenase can cause adverse drug reactions in patients receiving standard chemotherapy. This is another use of pharmacogenomics.

Chapter 125 - UTP

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