Pyrimidine nucleotides play a central role in metabolism

Mammalian Pyrimidine Biosynthesis:

Fresh Insights into an Ancient Pathway

David R. Evans and Hedeel I. Guy

Department of Biochemistry and Molecular Biology

Wayne State University School of Medicine

Detroit, MI 48201

Phone: (313) 577-1016

FAX: (313) 577-2765

Email: [email protected]

.

JBC Papers in Press. Published on April 19, 2004 as Manuscript R400007200

Copyright 2004 by The American Society for Biochemistry and Molecular Biology, Inc.

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Pyrimidine nucleotides play a critical role in cellular metabolism serving as activated

precursors of RNA and DNA, CDP-diacylglycerol phosphoglyceride for the assembly of cell

membranes and UDP-sugars for protein glycosylation and glycogen synthesis (1-3)*. In

addition, uridine nucleotides act via extracellular receptors to regulate a variety of physiological

processes (4)*. There are two routes to the synthesis of pyrimidines; nucleotides can be recycled

by the salvage pathways or synthesized de novo from small metabolites (Fig. 1). Most cells have

several specialized passive and active transporters (5*,6), that allow the reutilization of

preformed pyrimidine nucleosides and bases.

The relative contribution of the de novo and salvage pathways depends on cell type and

developmental stage. In general, the activity of the de novo pathway is low in resting or fully

differentiated cells where the need for pyrimidines is largely satisfied by the salvage pathways

(7). In contrast, de novo pyrimidine biosynthesis is indispensable in proliferating cells in order

to meet the increased demand for nucleic acid precursors and other cellular components.

Consequently, the activity of the de novo pathway is subject to elaborate growth state dependent

control mechanisms. Pyrimidine biosynthesis is invariably up-regulated in tumors and

neoplastic cells (8)* and the pathway has been linked to the etiology or treatment of several other

disorders including AIDS (9), diabetes (10), and various autoimmune diseases (11) such as

rheumatoid arthritis. This review focuses on the structure and regulation of the pyrimidine

biosynthetic complexes and the interplay of the diverse control mechanisms operative in

mammalian cells.

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The multifunctional protein CAD

Since the pioneering discoveries of Jones, Hoogenraad and others in 1971 (1)*, the

physical association of the first three enzymes of the de novo pyrimidine pathway, carbamoyl

phosphate synthetase (CPSase), aspartate transcarbamoylase (ATCase) and dihydroorotase

(DHOase) has been documented in many animal cells. In 1977, Stark and his associates made

the remarkable discovery (12) that all three activities are carried on a single polypeptide. The

243 kDa CAD polypeptide associates to form hexamers and higher oligomers (13) so that the

mass of the complex exceeds 1.4 MDa or about half the size of the ribosome. The domain

structure (Fig. 2) has been mapped and the function of each domain has been assigned (14).

Carbamoyl Phosphate Synthetase: CPSase catalyzes the synthesis of carbamoyl phosphate

from glutamine, bicarbonate and two ATP molecules. Carbamoyl phosphate biosynthesis (Fig.

1, ) is a complex process involving four partial reactions (15,16)* catalyzed by the GLN,

CPS.A and CPS.B domains of the molecule (Fig. 2). The 40 kDa GLN domain is an

amidotransferase that generates ammonia by glutamine hydrolysis. The synthetase domain,

consists of two homologous 60 kDa halves, CPS.A and CPS.B thought to have arisen by an

ancestral gene duplication and fusion (17). These subdomains are functionally equivalent (18),

but assume specialized functions when fused together in the intact molecule (19,20). CPS.A

catalyzes the ATP-dependent activation of bicarbonate forming carboxy phosphate which then

reacts with ammonia to form carbamate. Carbamoyl phosphate is formed in a second ATP-

dependent phosphorylation on CPS.B. All of these intermediates are labile but are effectively

sequestered within the complex, passing between the active sites via narrow tunnels that snake

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through the interior of the molecule (21). CPSase catalyzes the rate limiting step in de novo

pyrimidine biosynthesis and, as discussed below, controls the flux through the pathway (1)*.

Aspartate Transcarbamoylase: The reaction of carbamoyl phosphate and aspartate to form

carbamoyl aspartate is catalyzed by ATCase (Fig. 1 ). A single CPSase in E. coli supplies

carbamoyl phosphate for both pyrimidine and arginine biosynthesis. Thus, the bacterial ATCase

catalyzes the first committed step in the pyrimidine pathway and is allosterically regulated (22)*.

In contrast, there are two CPSases in mammalian cells, CPSII, the CAD activity committed to

pyrimidine biosynthesis, and CPSI (23)*, an ammonia dependent enzyme that initiates urea

biosynthesis in the mitochondria. Since CAD CPSase catalyzes the first step in the pathway,

ATCase is unregulated and there is no counterpart of the regulatory chain found in E. coli

ATCase. The isolated CAD ATCase domain, from proteolytic digests (24) or expressed in E.

coli (25), is a homotrimer of 34 kDa subunits. Kinetic and modeling studies showed that the

mammalian domain shares a common catalytic mechanism, oligomeric structure and tertiary fold

with the E. coli ATCase catalytic subunit (24-27), including a composite active site comprised of

residues from adjacent subunits. Qiu and Davidson (27) showed that substitutions at the trimer

interface disrupts the CAD oligomeric structure suggesting that the trimeric ATCase interactions

are a crucial organizing element in the hexamer.

Dihydroorotase: The 46 kDa CAD DHOase domain, which catalyzes the reversible

condensation of carbamoyl aspartate to dihydroorotate (Fig. 1 ), is a zinc metalloenzyme.

While the active site of E. coli DHOase has two zinc ions and a carboxylysine that bridges the

metal centers (28), the mammalian DHOase domain probably belongs to a different subgroup of

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the amidohydrolase superfamily (29). An extensive phylogenetic analysis (30) classified

DHOases into two major classes thought to have arisen by an ancestral gene duplication. Type I

DHOases are the most ancient and include domains of multifunctional proteins, such as CAD,

subunits of multienzyme complexes and monofunctional enzymes. The type II enzymes, such as

E. coli DHOase, are a more recent evolutionary development, are smaller (38 kDa) and have

undergone appreciable changes in sequence. The isolated CAD DHOase domain (31-34) has

only one zinc atom and is larger than its bacterial counterpart consistent with its assignment as a

type I DHOase.

Dihydroorotate Dehydrogenase

Mammalian DHOdhase is a 43 kDa flavoprotein (FMN) localized in the mitochondria

that oxidizes dihydroorotate to orotate (Fig. 1 ). The electrons are transferred directly to the

respiratory chain via ubiquinone (Q) (1). Biochemical and microscopic studies (1,35) showed

that the enzyme is an integral membrane protein localized in the inner mitochondrial membrane

with the active site facing the inner membrane space (Fig. 3). Mitochondrial import (36) is

governed by an uncleaved bipartite sequence at the amino end of the polypeptide (Figs. 2, 3) that

consists of a mitochondrial targeting (MT) sequence and a membrane stop-transfer (MA)

sequence that anchors the protein to the inner membrane.

Structural studies showed that tuncated human DHOdhase (37), lacking the bipartite

sequence, consists of two domains (Fig. 2). The active site is located within the large catalytic

domain (CAT), while the small domain (QT) forms a tunnel that leads directly to the bound

FMN and provides access to ubiquinone. Orotate (oa) is completely buried on the distal side of

FMN so it is unlikely to enter via the same tunnel. Instead, a flexible loop moves out of position

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to accommodate substrate binding on the distal side of FMN. Antiproliferative agents such as

leflunomide, bind within the tunnel blocking access of ubiquinone to the active site.

The multifuctional protein UMP Synthase

UMP synthase is a bifunctional protein that catalyzes the last two steps of de novo

pyrimidine biosynthesis (Fig. 1) (1)*. The mammalian protein (Fig. 2) consists of a 24 kDa

orotate phosphoribosyltransferase (OPRTase) that catalyzes the transfer of PRPP to orotate

forming OMP (Fig. 1 ) and a 28 kDa orotidine-5’-phosphate decarboxylase (ODCase) that

decarboxylates OMP (Fig. 1 ) forming UMP, (38). Sedimentation analysis of mouse UMP

synthase (39) identified a monomeric and two dimeric species, 5.1S and 5.6S. The monomer

lacks ODCase activity and the 5.1S dimer is only partially active. The formation of the fully

active 5.6S dimer is induced by the binding of OMP or nucleotide analogs.

CTP Synthase

CTPSase (Fig. 1 ) catalyzes the ATP dependent transfer of the amide nitrogen of

glutamine to the C-4 position of UTP to form CTP (40)*. While there are no structural studies,

sequence analysis revealed that CTPSase consists of two domains (Fig. 2); an amidotransferase

domain (GLN) that hydrolyzes glutamine and an amidator (AMD) domain that catalyzes the

ATP dependent phosphorylation of UTP and its subsequent reaction with ammonia to form CTP.

This reaction is the rate limiting step in the formation of cytosine nucleotides and as such

represents another important control locus in pyrimidine biosynthesis.

Intracellular Location of the Pyrimidine Biosynthetic Enzymes

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Biochemical and microscopic studies showed that CAD is primarily cytosolic (35,41)

with a smaller fraction in the nucleus. In the cytosolic compartment, CAD and UMP synthase

are localized around and outside the mitocondria and CAD appears to be associated with the

cytoskeleton. Mitochondria are known to be anchored to the cytoskeletal network, so an

interesting possibility is that CAD binds to and translocates along the filament to the

mitochondria where DHOdhase is located (Fig. 4, path A). The physical association of CAD

with the mitochondria is an attractive idea since under physiological conditions, the equilibrium

strongly favors the formation of carbamoyl aspartate over dihydroorotate (42). Docking CAD

near the mitochondria may allow a more efficient capture of dihydroorotate by DHOdhase and

prevent the accumulation of carbamoyl aspartate in the cell.

The role of CAD in the nucleus was initially not given much credence (41). However,

Angeletti and Engler (43) subsequently found that CAD is associated with the nuclear matrix in

adenovirus infected cells, where it anchors the pTP protein at sites of active replication.

Moreover, the ura2 protein, a CAD homolog in Saccharomyces cerevisiae consisting of CPSase,

ATCase and an inactive DHOase domain, was found within the nucleus (44).

More recently (45), a plasmid encoding a CAD-GFP fusion protein has been transfected

into BHK cells making it possible to monitor the movement of CAD in live cells. These studies

showed that CAD is localized in the cytosol in resting cells or during the G1 phase of the cell

cycle, but that a substantial fraction, 30%, is translocated into the nucleus during the S phase

when the demand for pyrimidine nucleotides reaches a peak.

The localization of CAD in the nucleus near the site of RNA and DNA synthesis, might

seem to be an efficient arrangement, but the exclusive localization of DHOdhase within the

mitochondria undermines this argument. Dihydroorotate synthesized in the nucleus would have

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to exit, diffuse into the mitochondria where it is oxidized to orotate, and orotate would then

reenter the cytoplasm (Fig. 4, path B). This circuitous route would seem to negate the putative

functional efficiency of locating CAD in the nucleus. Thus, the role of nuclear CAD remains

uncertain, although it is possible that nucleocytoplasmic translocation plays a role in the

regulation of the pathway or that CAD has a moonlighting function, perhaps in DNA replication

or cell division, that is unrelated to pyrimidine biosynthesis.

Regulation of Pyrimidine Biosynthesis

The intracellular nucleotide pools are controlled within narrow limits in normal, resting

cells but expand three to four fold in tumor cells (3)*. Even larger, 8-fold, increases occur in

mitogen stimulated lymphocytes (7). The increased demand for nucleotides is satisfied in large

part by up-regulation of de novo pyrimidine biosynthesis as a result of increased intracellular

enzyme levels and metabolic control mechanisms.

Gene Expression: CAD gene expression is controlled at both the transcriptional and

posttranscriptional level and is up-regulated when resting cells enter the proliferative phase (46-

48). Myc binding to an upstream E box was found to be responsible for the increase in CAD

gene transcription that occurs at the G1/S boundary as cells traverse the cycle (49,50). At the

protein level, CAD is rapidly degraded at the onset of apoptosis by caspase cleavage (51)*. Less

is known about the transcriptional regulation of the genes encoding the other pyrimidine

biosynthetic enzymes, but there are many studies that show that the intracellular concentrations

of these enzymes, and in some instances the mRNA, are appreciably higher in tumors and other

rapidly growing cells (8)*.

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The rate of cell growth probably sets the level of CAD and other enzymes and establishes

the basal rate of de novo pyrimidine biosynthesis, but the rapid changes in flux through the

pathway requires precise metabolic control exerted by allosteric effectors and the activity of

signaling cascades.

Metabolic Control: The CPSase activity of CAD is the major locus of control of de novo

pyrimidine biosynthesis (1). The enzyme is subject to feedback inhibition by the end product

UTP and is allosterically activated by PRPP. PRPP, a UMP synthase substrate (Fig. 1), is a

feedforward activator and helps coordinate pyrimidine and purine production since it is also a

substrate in the first step of purine biosynthesis. The allosteric effectors bind to a regulatory

subdomain (Fig. 2, B3) at the carboxyl end of CPS.B in CAD (52) and other CPSases (53,54).

Several studies (1)* suggest that allosteric regulation of CAD governs the rate of pyrimidine

biosynthesis in vivo. For example, expansion and depletion of the UTP and PRPP pools in

cultured cells, tissue slices and whole animals produce dramatic effects on the activity of the

pyrimidine biosynthetic pathway that correlate with the effects of these ligands on the CAD

CPSase activity. A nice illustration is provided by a recent analysis (55) showing that the

suppressor of black mutation in Drosophila disables UTP inhibition of CAD allowing the

pyrimidine pools to expand, thus providing the β-alanine precursors needed for normal cuticle

pigmentation.

Carrey made the important discovery (56) that cAMP-dependent protein kinase A (PKA)

phosphorylates purified CAD at two sites (Fig. 2). Modification of Ser1406 in the regulatory

subdomain (Fig. 2) abolishes UTP inhibition (56) and appreciably decreases the affinity of the

enzyme for PRPP (57). The loss of feedback inhibition would be expected to stimulate

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pyrimidine biosynthesis and allow unchecked expansion of the nucleotide pools needed for cell

proliferation, however, this effect would be partially offset by the reduction of PRPP activation.

Further insight into the growth state dependent regulation of the pathway was provided

by the discovery (58) that epidermal growth factor (EGF) stimulation promotes the

phosphorylation, via the MAP kinase (Erk1/2) cascade, of a single residue, Thr456, in the

CPS.A (A1) domain of CAD (Fig. 2). This modification has no effect on any CAD activity, but

converts UTP from an inhibitor to a modest activator and appreciably stimulates PRPP

activation. Both changes in the allosteric transitions would be expected to increase the flux

through the pyrimidine pathway. Moreover, a specific Thr residue (Fig. 2, Thr1037) in the B1

subdomain was found to be autophosphorylated both in vivo and in vitro (59), resulting in a

selective increase in CPSase activity and modulation of the allosteric transitions.

CTP synthase activity regulates the relative size of the UTP and CTP pools and also

helps coordinate the production of pyrimidine and purine nucleotides. The enzyme is

allosterically activated by GTP and is inhibited by the end product CTP (Fig. 1). The yeast

enzyme is phosphorylated by both PKA and protein kinase C (PKC) (60). The phosphorylated

enzyme is a dimer but the binding of the substrates, UTP and ATP, induces the formation of the

active tetramer. In contrast, the dephosphorylated enzyme is an inactive dimer that does not

undergo nucleotide induced tetramerization. Less is known about the mammalian enzyme.

Human CTPSase is regulated by GTP and CTP (61), but whether or not it is also controlled by

phosphorylation is unknown.

Growth State Dependent Regulation of the de novo pathway: A low basal level of

pyrimidine biosynthesis is needed to sustain resting cells. The activity of the pathway increases

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8-fold when BHK cells enter the exponential growth phase and then drops precipitously to basal

levels as the culture becomes confluent (62). The transition to exponential growth is associated

with a large increase in MAP kinase activity and the phosphorylation of the CAD MAP kinase

site (Thr456). As a result, UTP inhibition of CAD is abolished and PRPP activation increases

21-fold, changes in the allosteric transitions that can account for the stimulation of pyrimidine

biosynthesis. As the cultures approach confluence and growth ceases, Thr456 is

dephosphorylated and there is a concomitant increase in PKA phosphorylation of CAD. The

response to PRPP rapidly decreases and the activity of the pyrimidine biosynthetic pathway is

down regulated. The sequential changes in CAD phosphorylation state coincide with the

upregulation of the pathway as the cells approach S phase and are reversed at the S/G2 boundary

as pyrimidine biosynthesis is down-regulated (45). The lack of down regulation of pyrimidine

biosynthesis in tumorigenic breast cancer cells, MCF7, has been attributed to the elevated MAP

kinase activity that leads to persistent phosphorylation of the CAD MAP kinase site and a

concomitant blockage of PKA phosphorylation of CAD (63).

These observations are consistent with a model (Fig. 5) in which the rate of pyrimidine

biosynthesis is constrained in resting cells by UTP inhibition. As the cells enter the cycle, the

MAP kinase (MK) mediated phosphorylation of CAD serves as a molecular switch uncoupling

feedback inhibition and allowing the nucleotide pools to expand. CAD activity is now primarily

controlled by PRPP which maximally activates during S phase as a result of the sequential MAP

kinase and PKA mediated phosphorylations.

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Conclusions

The steps involved in pyrimidine biosynthesis occur nearly universally in all organisms,

however, compared to the monofunctional bacterial proteins, the eukaryotic enzymes have a

more complex structural organization and a more sophisticated mode of control. The pathway is

subject to diverse regulatory mechanisms including allosteric inhibition and activation,

phosphorylation and perhaps changes in intracellular location. Deciphering the operation and

interplay of these controls in the cell remains a fascinating challenge.

References

1. Jones, M. E. (1980) Ann. Rev. Biochem. 49, 253-279

2. Evans, D. R. (1986) in Multidomain Proteins-Structure and Function, (Hardie, D. G. a.

C., J.R., ed), pp. 283-331, Elsevier Science Publishers

3. Traut, T. W. (1994) Mol. Cell. Biochem. 140, 1-22

4. Connolly, G. P., and Duley, J. A. (1999) Trends Pharmacol Sci 20, 218-25.

5. Cass, C. E., Young, J. D., and Baldwin, S. A. (1998) Biochem Cell Biol 76, 761-70.

6. Mascia, L., Turchi, G., Bemi, V., and Ipata, P. L. (2001) Biochim Biophys Acta 1524, 45-

50.

7. Fairbanks, L. D., Bofill, M., Ruckemann, K., and Simmonds, H. A. (1995) J Biol Chem

270, 29682-9.

8. Weber, G. (2001) Biochemistry (Moscow) 66, 1164-73.

9. Bofill, M., Fairbanks, L., Ruckemann, K., Lipman, M., and Simmonds, H. (1995) J. Biol.

Chem. 270, 29690-29697

by guest on April 8, 2018

http://ww

w.jbc.org/

Dow

nloaded from

13

10. Kunjara, S., Sochor, M., Ali, M., Drake, A., Greenbaum, A., and McLean, P. (1991)

Biochem. Med. Metab. Biol. 46, 215-225

11. Sanders, S., and Harisdangkul, V. (2002) Am. J. Med. Sci. 323, 190-193

12. Coleman, P., Suttle, D., and Stark, G. (1977) J. Biol. Chem. 252, 6379-6385

13. Lee, L., Kelly, R. E., Pastra-Landis, S. C., and Evans, D. R. (1985) Proc Natl Acad Sci U

S A 82, 6802-6

14. Kim, H., Kelly, R. E., and Evans, D. R. (1992) J. Biol. Chem. 267, 7177-84

15. Anderson, P. M. (1995) in Nitrogen Metabolism and Excretion (Walsh, P. J., and Wright,

P. A., eds), pp. 33-55, CRC Press

16. Evans, D. (2002) in Wiley Encyclopedia of Molecular Medicine Vol. 5, pp. 449-453,

John Wiley and Sons, Inc., New York

17. Nyunoya, H., Broglie, K. E., and Lusty, C. J. (1985) Proc Natl Acad Sci U S A 82, 2244-

6

18. Guy, H. I., and Evans, D. R. (1996) J. Biol. Chem. 272, 13762-13769

19. Britton, H. G., Rubio, V., and Grisolia, S. (1979) Eur J Biochem 102, 521-30

20. Post, L. E., Post, D. J., and Raushel, F. M. (1990) J Biol Chem 265, 7742-7

21. Thoden, J. B., Holden, H. M., Wesenberg, G., Raushel, F. M., and Rayment, I. (1997)

Biochemistry 36, 6305-6316

22. Lipscomb, W. N. (1994) Adv Enzymol Relat Areas Mol Biol 68, 67-151

23. Lusty, C. J. (1978) Eur J Biochem 85, 373-83

24. Grayson, D. R., and Evans, D. R. (1983) J Biol Chem 258, 4123-9

25. Maley, J. A., and Davidson, J. N. (1988) Mol Gen Genet 213, 278-84

26. Scully, J. L., and Evans, D. R. (1991) Proteins 9, 191-206

by guest on April 8, 2018

http://ww

w.jbc.org/

Dow

nloaded from

14

27. Qiu, Y., and Davidson, J. N. (2000) Proc Natl Acad Sci U S A 97, 97-102

28. Thoden, J. B., Phillips, G. N., Jr., Neal, T. M., Raushel, F. M., and Holden, H. M. (2001)

Biochemistry 40, 6989-97.

29. Holm, L., and Sander, C. (1997) Proteins 28, 72-82.

30. Fields, C., Brichta, D., Shephardson, M., Farinha, M., and O'Donovan, G. (1999) Paths to

Pyrimidines 7, 49-63

31. Kelly, R. E., Mally, M. I., and Evans, D. R. (1986) J Biol Chem 261, 6073-83

32. Musmanno, L. A., Maley, J. A., and Davidson, J. N. (1991) Gene 99, 211-6

33. Williams, N. K., Peide, Y., Seymour, K. K., Ralston, G. B., and Christopherson, R. I.

(1993) Protein Eng 6, 333-40

34. Zimmermann, B. H., and Evans, D. R. (1993) Biochemistry 32, 1519-27

35. Carrey, E. A., Dietz, C., Glubb, D. M., Loffler, M., Lucocq, J. M., and Watson, P. F.

(2002) Reproduction 123, 757-68.

36. Rawls, J., Knecht, W., Diekert, K., Lill, R., and Loffler, M. (2000) Eur J Biochem 267,

2079-87.

37. Liu, S., Neidhardt, E. A., Grossman, T. H., Ocain, T., and Clardy, J. (2000) Structure

Fold Des 8, 25-33.

38. Suttle, D. P., Bugg, B. Y., Winkler, J. K., and Kanalas, J. J. (1988) Proc Natl Acad Sci U

S A 85, 1754-8.

39. Traut, T. W., and Payne, R. C. (1980) Biochemistry 19, 6068-74.

40. Zalkin, H. (1985) Methods Enzymol 113, 282-7

41. Chaparian, M. G., and Evans, D. R. (1988) Faseb J 2, 2982-9

42. Christopherson, R. I., and Jones, M. E. (1980) J Biol Chem 255, 3358-70

by guest on April 8, 2018

http://ww

w.jbc.org/

Dow

nloaded from

15

43. Angeletti, P. C., and Engler, J. A. (1998) J Virol 72, 2896-904

44. Nagy, M., Le Gouar, M., Potier, S., Souciet, J. L., and Herve, G. (1989) J Biol Chem 264,

8366-74

45. Sigoillot, F., Berkowski, J., Sigoillot, S., Kotsis, D., and Guy, H. (2003) J. Biol. Chem.

278, 3403-3409

46. Rao, G. N., and Davidson, J. N. (1988) Dna 7, 423-32

47. Rao, G. N., Church, R. L., and Davidson, J. N. (1988) FEBS Lett 232, 238-42

48. Rao, G. N., and Church, R. L. (1988) Exp Cell Res 178, 449-56

49. Boyd, K. E., and Farnham, P. J. (1997) Mol Cell Biol 17, 2529-37.

50. Khan, S., Abdelrahim, M., Samudio, I., and Safe, S. (2003) Endocrinology 144, 2325-35.

51. Huang, M., and Graves, L. M. (2003) Cell Mol Life Sci 60, 321-36.

52. Liu, X., Guy, H. I., and Evans, D. R. (1994) J Biol Chem 269, 27747-27755

53. McCudden, C. R., and Powers-Lee, S. G. (1996) J. Biol. Chem. 271, 18285-18294

54. Bueso, J., Cervera, J., Fresquet, V., Marina, A., Lusty, C. J., and Rubio, V. (1999)

Biochemistry 38, 3910-7.

55. Simmons, A. J., Rawls, J. M., Piskur, J., and Davidson, J. N. (1999) J Mol Biol 287, 277-

85.

56. Carrey, E. A., Campbell, D. G., and Hardie, D. G. (1985) Embo J 4, 3735-42

57. Sahay, N., Guy, H. I., Xin, L., and Evans, D. R. (1998) J. Biol. Chem. 273, 31195-202

58. Graves, L. M., Guy, H. I., Kozlowski, P., Huang, M., Lazarowski, E., Pope, R. M.,

Collins, M. A., Dahlstrand, E. N., Earp III, H. S., and Evans, D. R. (2000) Nature 403,

328-332

59. Sigoillot, F. D., Evans, D. R., and Guy, H. I. (2002) J Biol Chem 277, 24809-17.

by guest on April 8, 2018

http://ww

w.jbc.org/

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nloaded from

16

60. Park, T. S., DJ, O. B., and Carman, G. M. (2003) J Biol Chem 278, 20785-94.

61. Van Kuilenburg, A. B., Elzinga, L., and Van Gennip, A. H. (1998) Adv Exp Med Biol

431, 255-8.

62. Sigoillot, F. D., Evans, D. R., and Guy, H. I. (2002) J Biol Chem 277, 15745-51.

63. Sigoillot , F., Sigoillot, S, and Guy, H. (2004) Int. J. Cancer 109, 491-8.

FOOTNOTES

*Review article. The references cited are by no means exhaustive and include review articles

and when possible only the most recent references. The reader is referred to the citations within

these sources for a more exhaustive bibliography.

bAbbreviations: AMD, amidator domain; ATCase, aspartate transcarbamoylase; casp;

carbamoyl aspartate; auto, CAD autophosphorylation site; CAD, multifunction protein that

initiates and regulates de novo pyrimidine biosynthesis, CPSase, carbamoyl phosphate

synthetase, CAT, catalytic domain; CTPSase, CTP Synthase; dho, dihydroorotate; DHOase,

dihydroorotase; DHOdhase, DHO dehydrogenase; gln, glutamine; GLN, glutamine

amidotransferase domain; GFP, green fluorescent protein; MA, membrane anchoring sequence;

MAPK, Erk 1/2 MAP kinase; MT, mitochondrial targeting sequence; ODC, orotidine-5’-

phosphate decarboxylase; oa, orotate; OPRTase, orotate phosphoribosyltransferase; QT,

DHOdhase tunnel domain; PKA, protein kinase A; PRPP, phosphoribosyl 5’-pyrophosphate;

UMP synthase, bifunctional protein with OPRT and ODC activities.

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Figure Legends

Figure 1. The de novo pyrimidine biosynthetic pathway, steps 1-6, and the subsequent

formation of CTP. UTP is formed from UMP by sequential reactions catalyzed by UMP/CMP

kinase ( ) and nucleoside diphosphate kinase ( ). The other enzymes are described in the text.

Loci of allosteric activation (+) and inhibition (-) are also shown.

Figure 2. The domain structure of the multifunctional proteins and enzymes that catalyze

pyrimidine biosynthesis. The region of CAD that binds UTP and PRPP, the CAD

autophosphorylation site (auto) and the CAD sites phosphorylated by MAP kinase (MAPK) and

PKA are indicated.

Figure 3. Schematic representation of DHOdhase bound to the inner mitochondrial membrane.

Figure 4. Intracellular distribution of CAD (blue spheres), DHOdhase (yellow spheres) and

UMP synthase (red spheres) and putative interactions with the nucleus, mitochondria and

cytoskeleton. The paths of CAD, both translocation along the filament (A) or entry into the

nucleus (B) is shown.

Figure 5. The cell cycle dependent regulation of pyrimidine biosynthesis. In proliferating cells,

the sensitivity of CAD to the activator PRPP sequentially increases (↑) and decreases (↓).

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18

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orotate OMP UMP UDP UTP CTP

HCO3-

2 ATPGln

carbamoylphosphate

carbamoylaspartate dihyroorotate

aspartate

ATPGln

PRPP+ -

-GTP

+

Figure 1

GLN CPS.A CPS.B DHO ATC

1 365 379 527 785 927 1060 1308 1456 1789 1918 2225

A1 A2 A3 B1 B2 B3

MAPK UTPPRPP

PKA1 PKA2(Thr456) (Ser1406) (Ser1859)

auto(Thr1037)

CAD

1 214 222 480

UMP Synthase

OPRT ODC

CTP Synthase

GLNAMD

1 300 540

DHOdhase

1 14 33 68 78 396

CATMT MA QT

Figure 2

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Figure 3

innermembrane

space

oa

FMN

innermembrane

outermembrane

dho mitotargetingsequence

(MT)

stoptransfer

sequence(MA)

tunneldomain

(QT)

flexibleloop

QQH2

CAD

DHOdhasecatalytic domain

(CAT)

UMP

UMPS

Figure 4

dho

dhooa

UMPUMP

HCO3-2ATPGln

HCO3-2ATPGln

mitochondria

nucleuscytoskeleton

UMPSynthase

CAD

DHOdhase

AB

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Figure 5

quiescentcells

G0 G1 S G2 M

DNA synthesis

pyrimidinebiosynthesis

proliferatingcells

PPP

P PP

MK

MK PKA

UTP PRPP↑ PRPP↓

ATP ADP ATP ADP

P

P

CAD

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David R. Evans and Hedeel I. GuyMammalian pyrimidine biosynthesis: Fresh insights into an ancient pathway

published online April 19, 2004J. Biol. Chem. 

  10.1074/jbc.R400007200Access the most updated version of this article at doi:

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