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Ribonucleotide reductase

ribonucleotide reductase M1 polypeptide
Symbol RRM1
Entrez 6240
HUGO 10451
OMIM 180410
RefSeq NM_001033
UniProt P23921
Other data
EC number
Locus Chr. 11 p15.5-15.4
ribonucleotide reductase M2 polypeptide
Symbol RRM2
Entrez 6241
HUGO 10452
OMIM 180390
RefSeq NM_001034
UniProt P31350
Other data
EC number
Locus Chr. 2 p25-p24

Ribonucleotide reductase (RNR) is an enzyme that controls the cellular concentration of deoxyribonucleotides. Biosynthesis begins with the building up of essential molecules that RNR processes in a catalyzed reaction to make deoxyribonucleotides. RNR assembles deoxyribonucleotides for the synthesis of DNA. The processes are identical in all living organisms. What makes RNR unique from other enzymes is the need for a free radical.

Deoxyribonucleotides are synthesised on the level of diphosphates. The substrates for RNR are ADP, GDP, CDP and UDP. dTDP is synthesised by another enzyme (thymidilate synthase) from dUMP.



The iron-dependent enzyme, ribonucleotide reductase (RNR), is essential for DNA synthesis. Class I RNR comprises RNR1 and RNR2 subunits, which can associate to form an active heterodimeric tetramer. Since the enzyme catalyses the de novo synthesis of deoxyribonucleotides (dNTPs), precursors to DNA synthesis, it is essential for cell proliferation.

Each RNR1 monomer consists of three domains: one mainly helical domain comprising the 220 N-terminal residues; a second large ten-stranded α/β structure [α-helix and β-sheet, Structural Classification of Proteins (SCOP)] comprising 480 residues; and a small five-stranded α/β structure comprising 70 residues (Jordan & Reichard, 1998). RNR2 contains a diferric iron center and a stable tyrosyl radical. In E. coli, the tyrosyl radical is located at position 122 (Y122) providing the stable radical for the Class I RNR2 subunits (Hogbom et al., 2001). In A. aegypti, this tyrosyl radical is located at position 184 (Y184) (Pham et al., 2002). The tyrosyl radical is deeply buried inside the protein in a hydrophobic environment, located close to the iron center that is used in the stabilization of a tyrosyl radical. The structure of two μ-oxo-linked irons is dominated by ligands that serve as iron binding sites: four carboxylates [aspartate (D146), glutamate (E177, E240, and E274)] and two histidines (H180 and H277) (Pham et al., 2002). Association occurs between the C-terminus of RNR2 and the C-terminus of RNR1 (Jordan & Reichard, 1998). Enzymatic activity is dependent on association of the RNR1 and RNR2 subunits. The active site consists of the active dithiol groups from the RNR1 as well as the diferric center and the tyrosyl radical from the RNR2 subunit.

Other residues of RNR2, such as aspartate (D273), tryptophan (W48), and tyrosine (Y356) further stabilize the active-site tyrosyl radical thus allowing electron transfer (Jordan & Reichard, 1998). These residues help in the transfer of the radical electron from tyrosine (Y122) of RNR2 to cysteine (C439) of RNR1. The electron transfer begins on RNR2 tyrosine (Y122) and continues in RNR2 to tryptophan (W48), which is separated from RNR1 tyrosine (Y731) by 2.5 nanometers. Electron transfer from RNR2 to RNR1 occurs via tyrosine (Y356 to Y731) and continues on through tyrosine (Y730) to cysteine (C439) in the active site (Chang et al., 2004). Site-directed mutations of the RNR primary structure indicate that all residues cited above participate in the long distance transfer of the free radical to the active site (Jordan & Reichard, 1998).

In Aedes aegypti mosquitoes, RNR2 retains most of the crucial amino acid residues, including aspartate (D64) and valine (V292 or V284), that are necessary in allosteric regulation; proline (P210 and P610), leucine (L453 and L473), and methionine (M603) residues that are located in the hydrophobic active site; cysteine (C225, C436 and C451) residues that are involved in removal of a hydrogen atom and transfer of the radical electron at the active site; cysteine (C225 and C436), asparagine (N434), and glutamate (E441) residues that bind the ribonucleotide substrate; tyrosine (Y723 and Y743) residues that dictate the radical transfer; and cysteine (C838 and C841) residues that are used in the regeneration of dithiol groups in the active site (Pham et al., 2002).


The enzyme ribonucleotide reductase (RNR) catalyzes the de novo synthesis of dNTPs (Nelson & Cox, 2000). Catalysis of ribonucleoside 5’-diphosphates (NDPs) involves a reduction at the 2’-carbon of ribose 5-phosphate to form the 2’-deoxy derivative-reduced 2’-deoxyribonucleoside 5’-diphosphates (dNDPs). This reduction is initiated with the generation of a free radical. Following a single reduction, RNR requires electrons donated from the dithiol groups of the protein thioredoxin. Regeneration of thioredoxin occurs when nicotinamide adenine dinucleotide phosphate (NADPH) provides two hydrogen atoms that are used to reduce the disulfide groups of thioredoxin.

Step 1 = an electron transfer on the RNR2 subunit activates a RNR1 cysteine residue in the active site with a free radical; Step 2 = the free radical forms a stable radical on C-3, and cysteine residue removes a hydrogen; Step 3 = a cation is formed on C-2 by transferring a hydrogen from a dithiol group and is stabilized by the radical, resulting in the loss of H2O from C-2; Step 4 = a hydrogen is transferred from the dithiol group to reduce the cation C-2; Step 5 = the C-3 radical is reduced by the hydrogen removed in step 2, and the tyrosyl radical is generated; Step 6 = redoxins transfer two hydrogen to the disulfide group that restores the original configuration.

Three classes of RNR have similar mechanisms for the reduction of NDPs, but differ in the domain that generates the free radical, the specific metal in the metalloprotein structure, and the electron donors. All classes use free-radical chemistry (Jordan & Reichard, 1998). Class I reductases use an iron center with ferrous to ferric conversion to generate a tyrosyl free radical. Reduction of NDP substrates occurs under aerobic conditions. Class I reductases are divided into IA and IB due to differences in regulation. Class IA reductases are distributed in eukaryotes, eubacteria, bacteriophages, and viruses. Class IB reductases are found in eubacteria. Class IB reductases can also use a radical generated with the stabilization of a binuclear manganese center. Class II reductases generate a free radical by mechanisms involving 5’-deoxyadenosyl cobalamin (coenzyme B12) and have a simpler structure than class I and class III reductases. Reduction of NDPs or ribonucleotide 5’-triphosphates (NTPs) occurs under either aerobic or anaerobic conditions. Class II reductases are distributed in archaebacteria, eubacteria, and bacteriophages. Class III reductases use a glycine radical generated with the help of an S-adenosyl methionine and an iron sulphur center. Reduction of NTPs is limited to anaerobic conditions. Class III reductases are distributed in archaebacteria, eubacteria, and bacteriophages (Jordan & Reichard, 1998 & Pham et al., 2002). Organisms are not limited to having one class of enzymes. For example, Escherichia coli have both class I and class III RNR.

Metabolic pathways

Several major pathways lead to the generation of precursors for the de novo synthesis of nucleotides. These pathways involve the generation of ribose 5-phosphate, carbon dioxide, amino acids and ammonia. Ribose 5-phosphate generation begins with a molecule of glucose that is oxidized via the pentose phosphate pathway. The pentose phosphate pathway produces NADPH for reducing power involved in the catalysis of NTPs to dNTPs, and to produce ribose 5-phosphate necessary for the synthesis of ribonucleotides. Carbon dioxide is always available for biosynthesis because its concentration in the blood is kept nearly constant via the bicarbonate buffer system. An important co-factor for ribonucleotide synthesis is tetrahydrofolate, which is the major mediator for carbon transfers. Its derivative, folate (a vitamin), cannot be synthesized in mammals. Many forms of tetrahydrofolate follow pathways that are interconnected. For ribonucleotide synthesis, the N10-formyl-tetrahydrofolate molecule is necessary for the transfer of formyl groups to the purine ring. Amino groups or ammonia are donated from the catabolism of amino acids beginning with a dietary protein molecule. The free ammonia is combined with glutamate by a reaction involving adenosine 5’-triphosphate (ATP) and the activity of glutamine synthetase, which produces a nontoxic molecule of glutamine that can be transported in the bloodstream. Glutamine synthetase is present in nearly all organisms and is allosterically regulated by end products of glutamine metabolism. During synthesis of purines, amino groups are removed from glutamine for purine rings. Purine ribonucleotides are attached to ribose 5-phosphate during assembly of intermediate inosinate (IMP) from precursors in the purine pathway, including glutamine, glycine, N10-formyl-tetrahydrofolate, bicarbonate, aspartate and ATP. Synthesis is catalyzed by large multienzyme complexes. Purine ribonucleotides are adenosine 5’-monophosphate (AMP) and guanosine 5’-monophosphate (GMP). AMP is formed from IMP by aspartate donating an amino group (leaving as fumarate) and guanosine 5’-triphosphate (GTP) providing a phosphate. GMP is formed by the oxidation of IMP at C-2 requiring NAD+. Following oxidation, glutamine donates an amino group (leaving as glutamate) then ATP provides a phosphate.

Pyrimidine ribonucleotides are formed from an orotate molecule that is assembled from aspartate to form the pyrimidine ring. Subsequently, orotate is attached to ribose 5-phosphate to yield orotidylate. These two steps are catalyzed by a large multienzyme complex (CAD). Pyrimidine ribonucleotides are cytidine 5’-monophosphate (CMP) and uridine 5’-monophosphate (UMP). Orotidylate is decarboxylated to form UMP. UMP and two ATPs are transferred by kinases to form uridine 5’-triphosphate (UTP). Cytidine 5’-triphosphate (CTP) is formed from UTP by glutamine donating an amino group (leaving as glutamate) and ATP providing a phosphate. In some species, ammonia can donate an amino group instead of glutamine. Generation of 2’-deoxythymidine 5’-monophosphate (dTMP) occurs by conversion of 2’-deoxyuridine 5’-monophosphate (dUMP). Thymidylate synthase catalyzes the reaction in which dTMP is formed from dUMP; to provide the carbon atom N5, N10-methylene-tetrahydrofolate is oxidized to 7, 8-dihydrofolate. Dihydrofolate reductase (DHFR) is an essential enzyme that regenerates tetrahydrofolate at the expense of NADPH. Ribonucleoside monophosphates (AMP, GMP, CMP, and UMP) are phosphorylated to ribonucleoside diphosphates for their particular base by specific kinases. Ribonucleoside diphosphates are phosphorylated a second time to ribonucleoside triphosphates by nucleoside diphosphate kinase, which is not specific for their base or for their 2’-carbon of ribose 5-phosphate and its 2’-deoxy derivative. The activity of nucleoside diphosphate kinase is sequential based on which class of RNR is used. These metabolic pathways generate the ribonucleotides (ATP, GTP, CTP, and UTP) that are precursors for dNTPs. Thus, RNR reduces the corresponding NTPs to dNTPs for DNA synthesis. Cellular concentration of dNTP is much lower than required for DNA replication, and RNR is essential for adequate precursors during DNA synthesis.

After RNR reduces NDP or NTP the enzyme becomes inactive because a disulfide bond is formed in the active site. An exchange reaction occurs that reduces the disulfide bond of RNR catalyzed by thioredoxin or glutaredoxin. RNR gains electrons on the active-site dithiol groups necessary for its activity.

Catalytic Reduction Mechanism

The mechanism that is currently accepted for the reduction of ribonucleotides to deoxyribonucleotides is depicted in the following scheme. The first step involves the abstraction of the 3’- H of substrate 1 by radical Cys439. Subsequently, the reaction involves the elimination of one water molecule from carbon C-2’ of the ribonucleotide, catalyzed by Cys225 and Glu441. In the third step there is a hydrogen atom transfer from Cys225 to carbon C-2’ of the 2’-ketyl radical 3, after previous proton transfer from Cys462 to Cys225. At the end of this step, a radical anionic disulfide bridge and the closed-shell ketone intermediate 4 are obtained. This intermediate has been identified during the conversion of several 2’-substituted substrate analogues, as well as with the natural substrate (Cerqueira, 2004) interacting with enzyme mutants. The next step is the oxidation of the anionic disulfide bridge, with concomitant reduction of the substrate, generating 5. The spin density shifts from the sulphur atoms to the C-3' atom of the substrate, with simultaneous proton transfer from Glu441 to carbon C-3'. The last step is the reverse of the first step and involves a hydrogen transfer from Cys439 to C-3’, regenerating the initial radical and resulting in the final product 6. Theoretical models of some steps of these mechanisms using the full model of the R1 protein can be found at the studies performed by Cerqueira et al. (Cerqueira, 2005 and Cerqueira, 2006)


Class I RNR comprises RNR1 and RNR2 subunits, which can associate to form a heterodimeric tetramer (Eklund et al., 1997). RNR1 contains both allosteric sites, mediating regulation of substrate specificity and activity (Uhlin and Eklund, 1994). Depending on the allosteric configuration, one of the four ribonucleotides binds to the active site.

Class I RNR is activated by binding ATP or inactivated by binding dATP to the activity site located on the RNR1 subunit. When the enzyme is activated, substrates are reduced if the corresponding effectors bind to the allosteric substrate specificity site. A = when dATP or ATP is bound at the allosteric site, the enzyme accepts UDP and CDP into the catalytic site; B = when dGTP is bound, ADP enters the catalytic site; C = when dTTP is bound, GDP enters the catalytic site. The substrates (ribonucleotides UDP, CDP, ADP, and GDP) are converted to dNTPs by a mechanism involving the generation of a free radical.

Regulation of RNR is designed to maintain balanced quantities of dNTPs. Binding of effector molecules either increases or decreases RNR activity. When ATP binds to the allosteric activity site, it activates RNR. In contrast, when dATP binds to this site, it deactivates RNR (Jordan & Reichard, 1998). In addition to controlling activity, the allosteric mechanism also regulates the substrate specificity and ensures the enzyme produces an equal amount of each dNTP for DNA synthesis (Jordan & Reichard, 1998). In all classes, binding of ATP or dATP to the allosteric site induces reduction of cytidine 5’-diphosphate (CDP) and uridine 5’-diphosphate (UDP); 2’-deoxyguanosine 5’-triphosphate (dGTP) induces reduction of adenosine 5’-diphosphate (ADP); and 2’-deoxythymidine 5’-triphosphate (dTTP) induces reduction of guanosine 5’-diphosphate (GDP) (Figure 1). Interestingly, class IB reductases are not inhibited by dATP because they lack approximately 50 N-terminal amino acids required for the allosteric activity site (Eliasson et al., 1996). Eukaryotic cells with class IA reductases have a mechanism of negative control to turn off synthesis of dNTPs as they accumulate. This mechanism protects the cell from toxic and mutagenic effects that can arise from the overproduction of dNTPs because changes in balanced dNTP pools lead to DNA damage and cell death (Kunz, 1988 & Meuth, 1989).

Inhibition of RNR1 and RNR2 structure

Generally Class I RNR inhibitors can be divided in three main groups: translation inhibitors, which unable the formation of the enzyme; dimerization inhibitors that prevent the complexation of the two RNR subunits (R1 and R2); and catalytic inhibitors that inactivate subunit R1 and/or subunit R2, leading to RNR inactivity (Cerqueira, 2005) .

Class I RNR can be inhibited by peptides similar to the C-terminus of RNR2. These peptides can compete with RNR2 for binding to RNR1, and as a result RNR1 does not form an enzymatically active complex with RNR2 (Hamann et al., 1998 & Climent et al., 1991). Although the C-terminus of RNR2 proteins is different across species, RNR2 can interact with RNR1 across species (Cosentino et al., 1991). When the mouse RNR2 C-terminus was replaced with the E. coli RNR2 C-terminal (7 or 33) amino acid residues, the chimeric RNR2 subunit still binds to mouse RNR1 subunits. However, they lack enzymatic activity due probably to the elimination of residues involved in the transfer of the free radical electron from the RNR2 to the RNR1 subunit (Hamann et al., 1998).

Small peptides can specifically inhibit the RNR2 subunits from binding with RNR1 when they share a significant similarity with the normal RNR2 C-terminus (Cooperman, 2003). This inhibition RNR2 binding to RNR1 has been tested successfully in herpes simplex virus (HSV) RNR. When a 7 amino acid oligomer (GAVVNDL) truncated from the C-terminus of the RNR2 subunit was used in competition assays, it prevented the normal RNR2 from forming an enzymatically active complex with RNR1 (Filatov et al., 1992). Other small peptide inhibitors similar to the RNR2 C-terminus have also been used successfully to inhibit HSV RNR enzymatic activity and thus HSV replication (Cohen et al., 1986). In mice, for the treatment of stromal keratitis and corneal neovascularization (HSV ocular disease), a small RNR2 C-terminal analog BILD 1263 has been reported to inhibit RNR and is effective in preventing these diseases (Brandt et al., 1996). In some cases, although treatment with small C-terminal analogs may not stop disease spreading, they can still help in healing. In the acyclovir-resistant HSV (PAAr5), a small peptide inhibitor BILD 1633 has been reported to be 5 to 10 times more potent than BILD 1263 against cutaneous PAAr5 infection (Duan et al., 1998). A combination therapy approach (BILD 1633 and acyclovir) is more effective to heal topical lesions in mice. These data suggest that small peptide inhibitors that compete with RNR2 for binding to RNR1 are useful in preventing the spread of HSV.?

The drugs Motexafin Gadolinium and hydroxyurea interferes with the action of this enzyme.


Brandt, C.R., Spencer, B., Imesch, P., Garneau, M., Déziel, R. (1996) Evaluation of a peptidomimetic ribonucleotide reductase inhibitor with a murine model of herpes simplex virus type 1 ocular disease. Antimicrob Agents and Chemother. 40(5): 1078-1084.

Cerqueira, NMFSA, Pereira, S, Fernandes, PA, Ramos, MJ.(2005) Overview of ribonucleotide reductase inhibitors: an appealing target in anti-tumour therapy. Curr Med Chem. 12 : 1283-1294.

Cerqueira, N. M. F. S. A., Fernandes, P. A., Eriksson, L. A., Ramos, M. J., (2004) Ribonucleotide activation by enzyme ribonucleotide reductase: understanding the role of the enzyme. Journal of Computational Chemistry, 25(16): 2031-7

Cerqueira, N. M. F. S. A., Fernandes, P. A., Eriksson, L. A., Ramos, M. J. (2006), Dehydration of Ribonucleotides catalysed by Ribonucleotide Reductase: The Role of the Enzyme. Biophys J 90(6): 2109-19.

Chang, M.C.Y., Yee, C.S., Stubbe, J., Nocera, D.G. (2004) Turning on ribonucleotide reductase by light-initiated amino acid radical generation. Proc. Natl. Acad. Sci. 101(18): 6882-6887.

Climent, I., Sjöberg, B.M., Huang, C.Y. (1991) Carboxyl-terminal peptides as probes for Escherichia coli ribonucleotide reductase subunit interaction: kinetic analysis of inhibition studies. Biochemistry. 30(21): 5164-5171.

Cohen, E.A., Gaudreau, P., Brazeau, P., Langelier, Y. (1986) Specific inhibition of herpes virus ribonucleotide reductase by a nanapeptide derived from the carboxy terminus of subunit 2. Nature. 321(6068): 441-443.

Cosentino, G., Lavallee, P., Rakhit, S., Plante, R., Gaudette, Y., Lawetz, C., Whitehead, P.W., Duceppe, J.S., Lepine-Frenette, C., Dansereau, N. (1991) Specific inhibition of ribonucleotide reductases by peptides corresponding to the C-terminal of their second subunit. Biochem. Cell Biol. 69(1): 79-83.

Cooperman, B.S. (2003) Oligopeptide inhibition of Class I ribonucleotide reductases. Biopolymers (Peptide Science). 71(2): 117-131.

Duan, J., Liuzzi, M., Paris, W., Lambert, M., Lawetz, C., Moss, N., Jaramillo, J., Gauthier, J., Déziel, R., Cordingley, M.G. (1998) Antiviral activity of a selective ribonucleotide reductase inhibitor against acyclovir-resistant herpes simplex virus type 1 in vivo. Antimicrob. Agents and Chemother. 42(7): 1629-1635.

Eklund, H., Eriksson, M., Uhlin, U., Nordlund, P., Logan, D. (1997) Ribonucleotide reductase—structural studies of a radical enzyme. Biol. Chem. 378(8): 821-825

Eliasson, R., Pontis, E., Jordan, A., Reichard, P. (1996) Allosteric regulation of the third ribonucleotide reductase (NrdEF enzyme) from enterobacteriaceae. J. Biol. Chem. 271(43): 26582-26587.

Filatov, D., Ingemarson, R., Gräslund, A., Thelander, L. (1992) The role of herpes simplex virus ribonucleotide reductase small subunit carboxyl terminus in subunit interaction and formation of iron-tyrosyl center structure. J. Biol. Chem. 267(22): 15816-15822.

Hamann, C.S., Lentainge, S., Li, L-S., Salem, J.S., Yang, F-D., Cooperman, B.S. (1998) Chimeric small subunit inhibitors of mammalian ribonucleotide reductase: a dual function for the R2 C-terminus? Protein Eng. 11(3): 219-224.

Hashemy, S. I., Ungerstedt, J. S., Zahedi Avval, F., and Holmgren, A. (2006). Motexafin gadolinium, a tumor-selective drug targeting thioredoxin reductase and ribonucleotide reductase. J Biol Chem 281, 10691-10697.

Hogbom, M., Andersson, M.E., Nordlund, P. (2001) Crystal structures of oxidized dinuclear manganese centres in Mn-substituted class I ribonucleotide reductase from Escherichia coli: carboxylate shifts with implications for O2 activation and radical generation. J. Biol. Inorg. Chem. 6(3): 315-323.

Jordan, A., Reichard P. (1998) Ribonucleotide reductases. Annu. Rev. Biochem. 67(1): 71-98.

Kunz, BA. (1988) Mutagenesis and deoxyribonucleotide pool imbalance. Mutat. Res. 200(1-2): 133-147.

Meuth, M. (1989) The molecular basis of mutations induced by deoxyribonucleoside triphosphate pool imbalances in mammalian cells. Exp. Cell Res. 181(2): 305-316.

Nelson, D. and Cox, M. (2000) Lehninger principles of biochemistry. Worth Publishers, New York.

Pham, D.Q.-D., Blachuta, B.J., Nichol, H., Winzerling, J.J. (2002) Ribonucleotide reductase subunits from the yellow fever mosquito, Aedes aegypti: cloning and expression. Insect Biochem. Mol Biol. 32(9): 1037-1044.

Rodionova, DA. and Gelfand MS. (2005). "Identification of a bacterial regulatory system for ribonucleotide reductases by phylogenetic profiling". Trends in Genetics 21 (7): 385–389. doi:10.1016/j.tig.2005.05.011.

Uhlin, U., Eklund, H. (1994) Structure of ribonucleotide reductase protein R1. Nature. 370(6490): 533-539.

External links

  • MeSH Ribonucleotide+reductases
This article is licensed under the GNU Free Documentation License. It uses material from the Wikipedia article "Ribonucleotide_reductase". A list of authors is available in Wikipedia.
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