Uracil in DNA – occurrence, consequences and repair


Uracil in DNA results from deamination of cytosine, resulting in mutagenic U : G mispairs, and misincorporation of dUMP, which gives a less harmful U : A pair. At least four different human DNA glycosylases may remove uracil and thus generate an abasic site, which is itself cytotoxic and potentially mutagenic. These enzymes are UNG, SMUG1, TDG and MBD4. The base excision repair process is completed either by a short patch- or long patch pathway, which largely use different proteins. UNG2 is a major nuclear uracil-DNA glycosylase central in removal of misincorporated dUMP in replication foci, but recent evidence also indicates an important role in repair of U : G mispairs and possibly U in single-stranded DNA. SMUG1 has broader specificity than UNG2 and may serve as a relatively efficient backup for UNG in repair of U : G mismatches and single-stranded DNA. TDG and MBD4 may have specialized roles in the repair of U and T in mismatches in CpG contexts. Recently, a role for UNG2, together with activation induced deaminase (AID) which generates uracil, has been demonstrated in immunoglobulin diversification. Studies are now underway to examine whether mice deficient in Ung develop lymphoproliferative malignancies and have a different life span.


The most common mutation in human cells is C→T transitions and this type of mutation is also found very frequently in human tumours. A major fraction of these transitions occurs in a CpG context and they have therefore been suggested to result from deamination of 5-methylcytosine (5-meC) to thymine, or cytosine to uracil. DNA replication would then insert A in the place of G (reviewed by Waters and Swann, 2000). However, C→T transitions are common in other sequences as well. Mismatches caused by deamination (T : G or U : G) are generally repaired by the base excision repair (BER) pathway. The BER pathway probably repairs the largest fraction of DNA damage, but its significance to human health remains unclear. The present review will focus on the repair of one of the most common lesions in DNA, uracil. However, since several of the known uracil-DNA glycosylases (UDGs) are not strictly uracil-specific, the repair of some uracil analogues (Figure 1), namely 5-hydroxymethyluracil (5-hmU), 3,N4-ethenocytosine (εC) and 5-fluorouracil (5-FU) will also be discussed. In addition, repair of T : G mismatches by DNA glycosylases will be addressed. It has become clear in recent years that BER for removal of uracil from DNA involves several DNA glycosylases, which may have different functional roles. The steps downstream of the initial glycosylase step may also differ, depending on origin and localisation of uracil, the specific DNA glycosylase involved, and timing in the cell cycle. In this review the emphasis will be on the enzymes that initiate the repair process, the DNA glycosylases.

Figure 1

Pairs and mispairs of uracil or uracil analogs with guanine or adenine. Ma=major grove; Mi=minor groove. Red arrows indicate how bases in mispairs have moved relative to normal base pairs

Uracil in DNA results from deamination of cytosine to uracil, creating a premutagenic U : G mispair, or from misincorporation of dUMP instead of dTMP during replication, creating a U : A pair (reviewed in Krokan et al., 1997). The inappropriate uracil is recognised by a UDG activity, which cleaves the N-glycosidic bond and leaves an abasic site in the DNA. The first UDG was discovered in Escherichia coli in 1974 in a search for activities that would repair uracil in DNA. This also represented the discovery of the BER pathway. The activity that was discovered released uracil as a free base, leaving an intact abasic site in DNA (Lindahl, 1974). Ung from the ung-gene later proved to be a representative of a highly conserved family of UDGs present in most living organisms examined (Olsen et al., 1989). However, mammalian cells contain at least three additional human glycosylases (TDG, SMUG1 and MBD4), that have the capacity to remove uracil from DNA (reviewed in Krokan et al., 2001). UNG and SMUG1 prefer single-stranded DNA as substrate, but also remove U from double-stranded DNA. In contrast, TDG and MBD4 are strictly specific for double-stranded DNA, and have very low turnover numbers (Table 1). The functional roles of the different enzymes are only starting to become understood. It seems clear that UNG has a central function in removal of U from misincorporated dUMP-residues (Nilsen et al., 2000; Otterlei et al., 1999). SMUG1 has tentatively been proposed to have a specific role in removal of deaminated cytosine residues (Nilsen et al., 2001), but this role needs more experimental confirmation. The roles of TDG (Hardeland et al., 2001) and MBD4 (Bellacosa, 2001) may be limited to repair of mismatched uracil, thymine and some damaged pyrimidines in double-stranded DNA, particularly in CpG and 5-methylCpG contexts. In addition, MBD4 apparently cross-talks with the mismatch repair (MMR) system through an interaction with MMR protein MLH1 (reviewed in Bellacosa, 2001). In the present paper, we focus on central and recent developments in the field. For more comprehensive overviews, the reader should consult previous reviews (Friedberg et al., 1995).

Table 1 Substrates of uracil-releasing enzymes

Generation and consequences of uracil in DNA

The chemical instability of cytosine in DNA (Shapiro, 1980) results in slow hydrolytic deamination that generates premutagenic G : U mispairs. These may cause G : C to A : T transitions unless repaired prior to the next round of replication, because DNA polymerases efficiently incorporate A opposite U in the template. Subsequent removal of U and insertion of T generate the A : T transition. The number of cytosine deaminations has been calculated to be in the order of 60–500 per genome per day. The uncertainty depends on the average fraction of DNA present in single stranded form, since deamination is 200–300-fold faster from single stranded DNA than from double stranded DNA (reviewed in Lindahl, 1993; Krokan et al., 1997). Cytosine in cyclobutane dimers deaminates some 7–8 orders of magnitude faster than other cytosines (Krokan et al., 1997) and has been implicated in mutagenesis both in bacteria (Horsfall et al., 1997) and in human cells (Tu et al., 1998). Whether uracil in dimers is subject to repair is apparently not known. Enzymatic deamination of cytosine by (cytosine-5)-methyltransferase (MT) may happen as an aberrant process when the cellular content of S-adenosylmethionine (SAM) is low, or when MT is very high, as observed in several tumours (Krokan et al., 1997). The significance of this type of enzymatic deamination of cytosine remains unclear. However, activation-induced deaminase (AID) was recently suggested to directly target C : G base pairs in DNA. AID was initially described as a B-cell specific protein proposed to be involved in RNA editing (Muramatsu et al., 1999) and also potentiates changes in immunoglobulin gene DNA (Muramatsu et al., 2000). Interestingly, expression of AID in E. coli gave a mutator phenotype that yielded nucleotide transitions at dC : dG in a context-dependent manner, and the frequency of these mutations was enhanced in cells lacking Ung (Petersen-Mahrt et al., 2002). It was thus proposed that AID-triggered deamination of dC to dU and subsequent removal of U by UNG was involved in the first phase of hypermutation driving immunoglobulin gene diversification (Petersen-Mahrt et al., 2002). Furthermore, inhibition of chicken UNG by expression of Ugi in DT40 B-cells alters the pathway of immunoglobulin hypermutation from transversion dominance (due to replication over sites of base loss) to transition dominance, as expected if the deaminated C is copied in replication (di Noia and Neuberger, 2002). How such a mechanism is regulated and targeted specifically to immunoglobulin genes in mammalian cells remains to be clarified. The proposed function of AID and UNG in immunoglobulin gene diversification (Petersen-Mahrt et al., 2002) makes studies on possible B-cell abnormalities particularly relevant in an Ung-deficient mouse model. Such abnormalities would probably occur late in life since Ung-deficient mice do not show obvious signs of phenotypic changes in the first 12 months (Nilsen et al., 2000). Studies on the possible occurrence of lymphomas in this model are now underway. The mechanism is still unclear, but one possibility would be that the delayed processing of AID-induced U : G mispairs results in abnormal recombinations due to strand breaks induced by DNA glycosylases other than Ung. It was recently shown that the CEM15 gene, which is similar to the AID gene, is inhibited by the HIV-1 Vif (virion infectivity factor) protein. The activity of CEM15 is still unknown (Sheehy et al., 2002).

The other main source of uracil in DNA is by incorporation of dUMP instead of dTMP during DNA replication. dUTP is a normally occurring intermediate in nucleotide metabolism, but the level is kept very low due to an efficient dUTPase which prevents incorporation of dUMP. Although incorporation of dUMP into DNA was demonstrated both in bacteria (Tye et al., 1977; Wist et al., 1978) and isolated mammalian nuclei 25 years ago (Wist et al., 1978), the extent and significance remain unclear. However, Ung-deficient mice have several-fold increased content of uracil in their genome even though the capacity to remove incorporated dUMP is not completely abolished (Nilsen et al., 2000). In addition, inhibition of nuclear UNG2 by antibodies reduced the ability of the nuclei to remove misincorporated dUMP (Otterlei et al., 1999). Together, these results indicate that dUMP is misincorporated to a significant extent and that UNG-proteins have a central role in removal of uracil in a U : A context. The full physiological significance of misincorporated dUMP remains unclear.

Human uracil-DNA glycosylase (UNG) encoded by the UNG gene

Mitochondrial UNG1 and nuclear UNG2 are major uracil-DNA glycosylases in human cells (Slupphaug et al., 1995). UNG2 has a specific role in removal of misincorporated uracil in an immediate post-replicative process, (Otterlei et al., 1999; Nilsen et al., 2000). However, it may also have an important role in removal of deaminated cytosine residues (Kavli et al., 2002).

A human uracil-DNA glycosylase activity was reported in 1976 (Sekiguchi et al., 1976), and shown to have a function in removal of incorporated dUMP from nuclear DNA (Wist et al., 1978). cDNA for human UNG was cloned (Olsen et al., 1989) based upon information on the amino acid sequence of the N-terminal end of human placental UNG (Wittwer et al., 1989). The sequence encoded by the cDNA demonstrated that the major human UNG was highly conserved in herpes viruses, bacteria, yeast and man (Olsen et al., 1989). The human UNG gene spans approximately 13.8 kb and is located at 12q24.1 (Krokan et al., 2001). The gene encodes both the mitochondrial form UNG1 and nuclear form UNG2, which have a common catalytic domain, but different N-terminal sequences (Nilsen et al., 1997). Promoters PB and PA direct synthesis of mRNA for UNG1 and UNG2, respectively and they are differentially regulated, with UNG2 mRNA as well as enzyme activity peaking in the S phase (Haug et al., 1998). Exon 1A encodes 44 amino acids unique to UNG2 and is spliced into codon 35 of exon 1B. All of exon1B is used to form mRNA for the unique N-terminal end of UNG1, which therefore contains 35 N-terminal amino acids not found in UNG2. Amino acids downstream of position 44 in UNG2 and 35 in UNG1 are identical in the two forms. The N-terminal end of UNG1 forms a classical and very strong mitochondrial targeting signal (MTS), most of which is removed by mitochondrial processing peptidase upon entry in the mitochondria (Bharati et al., 1998). Complete nuclear targeting requires approximately 100 of the UNG2 N-terminal amino acids, although the unique N-terminal 44 amino acids contain the most important residues in the nuclear localization signal (NLS). When fused to the N-terminal end of UNG2, the MTS overrides the NLS and directs mitochondrial targeting (Otterlei et al., 1998). The N-terminal 90 amino acids of UNG2 are readily lost due to proteolysis by cellular proteases during purification, but the compact and fully active catalytic domain is even resistant to Proteinase K treatment. The N-terminal part of UNG2 therefore probably forms an independent structural domain, or may be unstructured. UNG2 is located in the nucleoplasm and replication foci. The mechanism for entry of UNG2 into replication foci is not known, but it may be co-transported with proliferating cell nuclear antigen (PCNA), similar to the targeting of DNA ligase I (Rossi et al., 1999) and DNA pol ι (Haracska et al., 2001). The N-terminal part contains a classical motif (Q4xxLxxFF11) for binding of PCNA. In addition, the N-terminal part binds trimeric replication factor A (RPA) through a motif that overlaps with the PCNA-binding motif, and binds the 34 kD monomer of RPA by a second motif in residues 73–84. These interactions take place in replication foci (Otterlei et al., 1999). The interactions of UNG2 with PCNA and RPA, both putatively required for long patch BER as well as replication, and the presence of several other replication factors used in long patch repair, suggest that UNG2 is engaged in long patch repair in replication foci. However, UNG2 is also present in the nucleoplasm and may participate in repair of U : G mispairs resulting from cytosine deamination (Kavli et al., 2002). The specific role of UNG2 in removal of misincorporated uracil was demonstrated by the inhibition of immediate post-replicative removal of incorporated uracil in isolated nuclei by neutralizing anti-UNG antibodies (Otterlei et al., 1999), and by the slow removal of incorporated uracil in nuclei from Ung−/− mice (Nilsen et al., 2000). Incorporated uracil was eventually removed, however, indicating the presence in nuclei of a less efficient back-up system for removal of uracil (Nilsen et al., 2000). UNG2 is ideally suited for removal of misincorporated uracil close to the rapidly moving replication fork because it has a turnover number (600–1000 per min) which is orders of magnitude higher than that of other uracil-DNA glycosylases. In Ung−/− cells the steady state level of uracil in genomic DNA is approximately 2000 per cell, while it was below detection level, and at least 10-fold lower, in wild-type cells (Nilsen et al., 2000). Surprisingly, the spontaneous mutation level in Ung−/− mice was increased only 1.3-fold (thymus) and 1.4-fold (spleen) as compared with wild type. This is likely due to the presence of back-up activities, among which SMUG1 may have a dominant role (Nilsen et al., 2000, 2001). While these results are seemingly at odds with previous results in bacteria and yeast (reviewed in Krokan et al., 1997), a direct comparison between rapidly growing microorganisms and a mixture of cells in complex multicellular organisms may not be entirely informative. It is possible that mutation rates in a homogenous fraction of rapidly growing cells would be different. In fact, inhibition of UNG2 in human glioma cells by expression of the protein inhibitor Ugi resulted in a threefold increase in spontaneous mutation rates in a shuttle vector, which is comparable to the average mutation rates in microorganisms (Radany et al., 2000).

Several lines of evidence indicate that human UNG may also play a role in viral infection. Several DNA viruses such as the poxviruses and the herpesviruses encode their own UDG, whereas RNA viruses such as retroviruses do not. It could, however, be hypothesized that the latter utilizes cellular UNG to avoid misincorporation of uracil during synthesis of the replicative DNA intermediate. Interestingly, the HIV-1 proteins Vpr and integrase IN have both been shown to bind UNG (Bouhamdan et al., 1996; Willetts et al., 1999). Furthermore, human UNG was efficiently incorporated into wild-type HIV-1 virions. IN-defective viruses fail to incorporate UNG, indicating that the integrase is required for packaging of UNG into virions (Willetts et al., 1999). These findings indicate that the presence of UNG is important near the site of nascent viral DNA synthesis. Alternatively, the association of viral proteins with UNG might be important for their nuclear translocation. Further studies of the association between UNG and viral proteins may provide important insight into the biology of HIV-1 and other viruses.

The catalytic domain of UNG proteins

Our understanding of the individual molecular steps involved in enzymatic removal of uracil by the Ung proteins has largely evolved from analysis of the human, E. coli and the Herpes simplex virus type-1 UDGs. Crystal structures of HSV-1 (Savva et al., 1995), human (Slupphaug et al., 1996) and E. coli (Xiao et al., 1999) enzyme-ligand complexes provided decisive information on both the substrate recognition and the catalytic mechanism. The three enzymes are structurally essentially identical, and substrate binding occurs in a highly conserved pocket providing shape and electrostatic complementarity to uracil and which is too narrow to accommodate purines. Numbering of amino acids in human UNG below is with reference to UNG1 (Mol et al., 1995). UNG2 is nine amino acids longer at the unique N-terminal end (reviewed in Krokan et al., 2001). Selectivity against other pyrimidines is provided by human UNG Y147 (Y90 in HSV-1 and Y66 in E. coli), which exclude bulky 5-substitutions (such as thymine), and N204 (N147 in HSV-1 and N123 in E. coli) which specifically hydrogen bonds to N3 and O4 of uracil. Correct orientation of the latter amide group is fixed by a cluster of water molecules at the base of the uracil-binding pocket (Pearl, 2000). The contribution of both N204 as well as Y147 to specificity was also verified experimentally, as replacement of Asn204 with Asp in human UNG shifted the specificity of the mutant towards cytosine, while replacement of Y147 by the smaller Ala, Cys or Ser shifted the specificity towards thymine (Kavli et al., 1996).

An important conclusion from the crystal structures was that uracil within the helical context of DNA could not be accommodated within the buried uracil-binding pocket. Instead, uracil binding was likely mediated by ‘base flipping’. A conserved leucine (human L272), positioned directly above the uracil-binding pocket, was suggested as a candidate to assist the local melting of the DNA helix (Mol et al., 1995). Subsequently, base flipping by human UNG was directly visualized by cocrystallization of hUNG with a uracil-containing DNA duplex (Slupphaug et al., 1996). In this complex uracil, deoxyribose and 5′-phosphate were rotated 180° relative to their starting position in DNA, and the cleaved uracil was found in the specificity pocket. We thus suggested that the term ‘nucleotide flipping’ provided a better description of the process. Furthermore, compression of the backbone flanking uracil was for the first time implicated in catalysis, assisted by extensive conformational changes in the enzyme upon formation of the productive complex. This established a ‘push–pull’ hypothesis in which the L272 side chain penetrates into the minor groove expelling uracil (push), and complementary interactions from the specificity pocket facilitate productive binding (pull). The mechanistics of this was further refined by high resolution co-crystal structures of wild type and mutant UNG bound to U : A and U : G oligonucleotides (Parikh et al., 1998). In the L272A mutant structure the uracil had dissociated, and the enzyme rebound to the product, an extrahelically positioned AP-site. This implied that the extrahelical conformation could be achieved even in the absence of the insertion of a hydrophobic side chain (push). A mechanism was proposed in which serine-proline rich loops compress the phosphates flanking uracil by a ‘pinching’ mechanism and thereby stabilize the extrahelical conformation (Parikh et al., 1998) without contributing significant energy to the cleavage reaction itself (Werner et al., 2000). Such a ‘pinch–push–pull’ mechanism was also hypothesised for E. coli Ung (Stivers et al., 1999). More recent data, however, indicate that the function of the inserting leucine side chain may be more complex than merely pushing the uracil out of the double helix. When analysing kinetic parameters of E. coli Ung L191A against single-stranded substrates, a 15-fold reduction in kcat/Km was observed compared to the wild type (Handa et al., 2001; Jiang and Stivers, 2001), which would not be expected in the absence of stabilizing base pairing. A likely explanation for this could be that the leucine side chain rather functions as a ‘doorstop’ to prevent return of the flipped out uracil residue, as recently suggested by Wong et al. (2002). Using stopped-flow experiments of E. coli Ung and monitoring fluorescence of 2-aminopurine adjacent to uracil, they demonstrated that Leu191 insertion occurred after nucleotide-flipping but before excision, thus indicating a ‘pinch–pull–push’ rather than a ‘pinch–push–pull’ mechanism.

When considering the energy contribution of each discrete event above to the overall catalytic reaction, one should bear in mind that DNA is a very heterogeneous substrate. This is also reflected by the different efficiency whereby uracil is excized from different sequence contexts (Eftedal et al., 1993; Nilsen et al., 1995). Recently the sequence specificity was re-examined using both single-stranded and duplex DNA substrates (Bellamy and Baldwin, 2001), and the authors conclude that the observed variations were not due to stability of the uracil itself within the DNA structure. Rather, local structure perturbations could affect uracil recognition, e.g. by affecting the phosphate recognition by the serine pinch.

Uracil binding induces considerable conformational changes in UNG, bringing key residues in optimal distances to favour catalysis (Slupphaug et al., 1996). This is accompanied by large conformational strain induced upon the deoxyuridine (Parikh et al., 2000b; Werner and Stivers, 2000). U is both pulled relative to the sugar and rotated 90° around the N-glycosidic bond. This increases the orbital overlap and favours electron flow from O4′ of the deoxyribose to O2 of uracil. The developing negative charge at O2 is enzymatically stabilized by a neutral histidine (E. coli H187, human H268) (Drohat et al., 1999; Shroyer et al., 1999). Hydrolysis is completed by a weakly nucleophilic water molecule positioned directly below the deoxyribose C1′ that becomes the 1′-α-OH, and protonation of the uracil enolate anion N1 to give the more stable amide (Parikh et al., 2000b; Werner and Stivers, 2000). Moreover, recent quantum- and molecular-mechanical calculations indicate that negative phosphate charges in the substrate itself may repel the anionic leaving group, and thus make a major contribution to the catalytic rate (Dinner et al., 2001). The authors suggest that such substrate autocatalysis may emerge as a general feature of DNA glycosylases.

The observation that UNG had an higher affinity for the product AP-site than the actual substrate itself (Parikh et al., 1998), led to the assumption that subsequent to product release, UNG quickly rebound to the AP site to protect the cell until the AP site was conveyed to the next enzyme in the BER-pathway. Such rebinding has subsequently been observed for several DNA glycosylases (Vidal et al., 2001; Waters et al., 1999). Recent studies have, however, demonstrated that the human nuclear UNG2 does not bind AP-sites (Kavli et al., 2002), and the in vivo significance of AP-site shielding thus remains elusive, at least for UNG2.

Perhaps the least understood stage in the processing of uracil-DNA is how the glycosylases recognize these subtle lesions within vast stretches of DNA. This is further complicated by the fact that eukaryotic DNA is organized in complex nucleoprotein structures. In vitro, the UNG-proteins appear to function in both a processive and distributive fashion, depending on the salt concentration (Bennett et al., 1995; Slupphaug et al., 1995). When a uracil residue is encountered, the mechanism of initial recognition is not obvious. U : A base pairs only differ from T : A by the lack of a 5-methyl in the minor groove. U : G mispairs are somewhat more evident, as they exist in the wobble conformation with the uracil shifted towards the major groove leaving partially hydrophobic gaps within the base stack. Both U : A and U : G base pairs might conceivably be detected in the minor groove by the Leu272 loop (human) reading head. However, neither of these structural determinants appear necessary for base flipping to occur, since the UNG mutants TDG and CDG are able to recognize and cleave perfectly normal bases from the DNA (Kavli et al., 1996). Thus, the enzyme might instead flip every DNA base to probe against the specificity pocket. How the energetic cost of such a scanning mechanism is covered merits further investigation, however.

SMUG1 (Single strand-selective Monofunctional Uracil-DNA Glycosylase)

SMUG1 removes uracil, as well as 5-hydroxymethyluracil (5-hmeU), from single- and double stranded DNA and is proposed to have an important role in removal of uracil resulting from cytosine deamination (Nilsen et al., 2001). It also removes εC, although relatively inefficiently (Kavli et al., 2002). SMUG1 is not thought to have a role in removal of incorporated uracil and it does not accumulate in replication foci. SMUG1 was identified in extracts of Xenopus (xSMUG1) using a proteomics approach. This procedure comprised in vitro expression from a library of cDNA, and electrophoretic mobility shift upon binding of damage-recognising protein to DNA containing modified nucleotides designed to target the active site of the glycosylases. The human counterpart was identified from EST databases (Haushalter et al., 1999). Interestingly, the gene for human SMUG1, like TDG and UNG-proteins, is located in the long arm of chromosome 12, more specifically in position 12q13.1–14, centromeric of UNG and TDG which are located in positions 12q24.1 and 12q22–24.1, respectively (reviewed in Krokan et al., 2001). Thus, genes for three out of four uracil-removing activities are located on chromosome 12. These genes are thought to have evolved from a common ancestral gene by duplications and divergence and although the amino acid sequence conservation is <10%, sequence analysis suggests that the enzymes have retained a common fold (Pearl, 2000). The phylogeny of the uracil-DNA glycosylase genes will be discussed in more detail below.

The selectivity of SMUG1 for single-stranded DNA is more pronounced for the Xenopus enzyme than for the human enzyme, which in fact removes uracil relatively efficiently both from U : G and U : A contexts (Haushalter et al., 1999; Kavli et al., 2002). Thus, the term single-strand selective is not entirely appropriate for the human enzyme. The primary sequence of the SMUG1 proteins is very different from the UNG-proteins. Furthermore, the xSMUG1 activity was not inhibited by the peptide inhibitor Ugi that efficiently inhibits both prokaryotic and eukaryotic uracil-DNA glycosylases belonging to the Ung-family. Although hSMUG1 is weakly inhibited by Ugi (Kavli et al., 2002, data not shown), the DNA-binding region in SMUG1 is probably substantially different from the Ung-family of proteins. Human SMUG1 (hSMUG1) is a protein of predicted size 270 amino acids (30 kDa) and is 60% identical and 71% similar to the Xenopus enzyme. xSMUG1 acted upon U : G and U : A essentially without preference for either, but showed no enzymatic activity against T : G, in contrast to the T(U)-mismatch glycosylase TDG (Hardeland et al., 2001). Recently, it was found that SMUG1 is probably the major enzyme for removal of 5-hmeU (Boorstein et al., 1987) from DNA (Boorstein et al., 2001). 5-hmeU is a result of oxidation of the 5-methyl group of thymine in DNA due to ionizing radiation and other forms of oxidative stress (Faure et al., 1998). However, it may also be formed in a two-step reaction; first the methyl-group of 5-meC in CpG-contexts is oxidised to 5-hmeC, and subsequently this residue is deaminated to yield 5-hmeU (Cannon Carlson et al., 1989). The latter pathway might result in G : C to A : T transitions, but based upon information on rate constants for deamination and oxidation, the amount of 5-hmeU generated spontaneously by this scheme would be extremely low (Rusmintratip and Sowers, 2000).

In Ung−/− mice mSMUG1 is apparently the major uracil-DNA glycosylase activity, both in various organs and in mouse embryonal fibroblasts from knock-out mice (Nilsen et al., 2001). xSMUG1 has a low Km for U : G (0.035 μM), which would make it suitable for recognising U : G mismatches resulting from cytosine-deamination (Haushalter et al., 1999). Using a double-stranded DNA containing a single U : G mismatch at low substrate concentration to mimic the in vivo situation, mSMUG1 was found to be the major UDG activity in extracts of cells from Ung−/− mice. Even in extracts from wild-type mice, mSMUG1 contributed a substantial fraction of the total UDG-activity under these assay conditions. In addition, mSMUG1 activity is stimulated several-fold by APE1. These results would be compatible with mSMUG1 as the major enzyme responsible for removal of U from U : G mismatches (Nilsen et al., 2001). However, recent data from our laboratory on hSMUG1 as compared with hUNG2 are at variance with these results (Kavli et al., 2002). Thus, the Km values for hSMUG1 on double-stranded DNA containing U : G mismatches is not lower than the Km for hUNG2 tested under different conditions, including different concentrations of Mg2+. Mg2+, present in relatively high concentrations intracellularly, substantially enhances both hSMUG1 and hUNG2 activities. In the presence of 7.5 mM Mg2+, the Km values for hUNG2 and hSMUG1 against U : G are 0.4±0.1 μM and 1.3±0.2 μM, respectively. In fact, under all conditions tested with U : A, U : G and ssU, the Km for hUNG2 was either lower (most frequently), or similar to the Km values for hSMUG1. Furthermore, hUNG2 was also by far the most efficient enzyme in extracts from human tumor cells (CX-1, from a colorectal carcinoma) in removal of U from U : G mismatches, even when using the same oligonucleotide substrate described in the mSMUG1-study by Nilsen et al. (2001). Thus, at least in human cells both hUNG2 and hSMUG1 are candidates for removal of U from U : G mismatches, and if anything, hUNG2 is the major candidate with hSMUG1 as a back-up enzyme (Kavli et al., 2002). The situation may be different in mouse. hSMUG1 is a nuclear enzyme, but it is not cell cycle-regulated and in contrast to UNG2, it does not accumulate in replication foci (Kavli et al., 2002). This would agree with a function of hSMUG1 in removal of U from U : G mismatches independent of DNA replication. The efficiency of hSMUG1 in removal of 5hmU (kcat/Km) was 5hmU in ssDNA> 5hmeU : G>5hmU : A (Kavli et al., 2002).

In conclusion, hSMUG1 is a non-abundant enzyme present in the nucleoplasm. It efficiently removes U and 5hmUra from ssDNA and double-stranded DNA. It also inefficiently removes εC. The major function may be in removal of 5hmU and deaminated cytosines, although it may be less important than UNG2 in the latter process, at least in human cells.

Other activities for removal of 5-hmU from DNA

In addition to SMUG1, one well defined enzyme, TDG which has a strict requirement for dsDNA, removes 5-hmeU, but less efficiently than U and T (Rusmintratip and Sowers, 2000). In addition, an activity that removes 5-hmeU and that is different from SMUG1 and TDG has recently been partially purified. This enzyme has a strict requirement for double-stranded substrates (Baker et al., 2002). Thus, there are apparently at least three different human enzymatic activities for removal of 5-hmeU from DNA. While 5-hmeU-removing activity is relatively abundant in extracts from human nuclei, especially activity acting on 5-hmeU : G mismatches (Rusmintratip and Sowers, 2000), 5-hmeU-removing activity is apparently absent in bacteria and yeast (Boorstein et al., 1987). The 60-fold increased activity towards 5-hmeU : G over 5-hmeU : A may seem surprising, since by far the major route of formation of 5-hmeU is thought to be by oxidation of the methyl-group in thymine (Rusmintratip and Sowers, 2000). However, 5-hmeU : G mispairs may cause G : C to A : T transition mutations, while 5-hmeU : G may not, since 5-hmeU is not miscoding (Levy and Teebor, 1991), does not block polymerases (Vilpo and Vilpo, 1995), and does not perturb DNA structure (Mellac et al., 1993; Pasternack et al., 1996). These results may indicate that 5-hmeU : G mispairs may be generated more frequently than assumed, although decisive experimental evidence supporting this view is still missing.

TDG – a T(U) mismatch glycosylase that may have multiple functions

TDG removes T or U from mismatches in double-stranded DNA, with U : G being preferred over T : G. TDG also efficiently removes εC in mismatches, and removes 5hmU with lower efficiency. It is thought to have an important role in correcting mismatches resulting from deamination of C or 5-meC in CpG contexts, which are major sources of G→A transition mutations in man (Brown and Jiricny, 1987; Hardeland et al., 2001). However, the major physiological role of TDG remains elusive. Interestingly, TDG may also function as a transcription factor (Hardeland et al., 2001).

Identification of TDG and biochemical properties

TDG was discovered during a search for mammalian activities correcting T : G mismatches. After transfection of SV40 DNA containing this mismatch, C was found to be preferentially inserted in place of T in monkey cells (Brown and Jiricny, 1987). Later a protein of apparent size 55 kDa necessary for this repair was identified, and it was demonstrated to be a G : T mismatch glycosylase (TDG). Cloning of the cDNA revealed that the correct size of TDG was 46 kDa. The enzyme can also remove thymine from C : T and T : T mispairs. The order of efficiency is G : T>>C : T>T : T. It does not cleave the DNA backbone, and thus contains no lyase activity (Neddermann et al., 1996; Neddermann and Jiricny, 1993). However, TDG removes U mispaired to G with some 10-fold higher kcat than T from G : T mispairs (reviewed in Hardeland et al., 2001). The cyclic alkylation product εC is also a good substrate both for the bacterial homologue MUG and human TDG and removal of this lesion may be a major function of TDG (Barrett et al., 1999; Hang et al., 1998; Hardeland et al., 2001; Saparbaev and Laval, 1998). Surprisingly, TDG is able to process 5-fluorouracil (5-FU) in DNA with a higher efficiency than any other substrate that has been tested. The enzyme even removes 5-FU from single stranded DNA (Hardeland et al., 2001). This is unexpected since TDG is generally very double-strand-specific, and since the substrate recognition, as deduced from data on the homologous bacterial protein MUG, involves interactions with both strands (Barrett et al., 1999). Although different assay systems may influence the relative activities of TDG towards different substrates, 5-FU and εC appear to be the best substrates (Hardeland et al., 2001; Saparbaev and Laval, 1998), but in contrast to 5-FU, εC in single stranded oligonucleotides is not a substrate for TDG (Saparbaev and Laval, 1998). εC may result from exposure to environmental pollutants such as vinylchloride, but may also be caused by lipid peroxidation (reviewed in Bartsch and Nair, 2000). Since εC has lost the specific coding potential it is mutagenic, as well as cytotoxic, and causes both transitions and transversions (Basu et al., 1993; Moriya et al., 1994). It has been reported that TDG may also remove 5-meC from CpG contexts (Zhu et al., 2000b), but this has not been verified in a separate study (Hardeland et al., 2001).

In the cases described above, the substrate is a base that has been modified. However, TDG is also able to remove T from T : O6meG mismatches arising from miscoding O6meG residues in the template (reviewed in Hardeland et al., 2001). Repair of O6meG in mispairs by O6methylG-DNA methyltransferase would result in a G : T mispair that could either be repaired by TDG or the mismatch repair system. O6meG in the template may direct incorporation of either C or T. Thus, TDG may remove T and give the BER system a second chance to incorporate C opposite of the miscoding O6meG-residue.

Unlike UNG-proteins, isolated TDG has an extremely low catalytic turnover number. The rate-limiting step in the mechanism in vitro is the release of the product (e.g. G : AP-site) due to rigid hydrogen bonding interactions between the enzyme and the Watson-Crick face of the guanine opposite the AP-site (Hardeland et al., 2000). This shields the cytotoxic AP-site until the next enzyme in the pathway, AP-endonuclease APE1 (also called HAP1) enters. No evidence for direct interaction between TDG and APE1 has been found. Therefore, APE1 most likely displaces TDG because it has even higher affinity for the AP-site than TDG, and in addition APE1 is more abundant. In agreement with this, APE1 strongly stimulates the activity of TDG (Waters et al., 1999). Recent findings indicate, however, that the AP-site binding capacity of TDG might be subject to regulation. Hardeland et al. (2002) demonstrated that a substantial fraction of cellular TDG is covalently modified by the ubiquitin-like proteins SUMO-1 and SUMO-2/3. Furthermore, in vitro sumoylation of TDG dramatically reduced its DNA-substrate and AP-site affinity. This increased the enzymatic turnover of G : U substrates, with a concomitant loss of G : T-processing activity. The authors propose that TDG binds its substrate in the unmodified state, and that subsequent to catalysis, sumoylation allows detachment from the product AP-site. Interestingly, a sumoylation consensus site is also present in MBD4 (MED1), but the functional implications of this remain to be established.

Structural aspects

UNG and TDG have <10% identical amino acid residues, and would therefore not immediately be considered homologous proteins. However, they appear to have very similar structures. The structure of the bacterial homologue MUG has been solved (Barrett et al., 1998, 1999; Pearl, 2000). MUG is closely related (37% identity) to human TDG, and their structures must be similar. Crystal structures of MUG and UNG reveal a striking homology, containing similarly positioned α-helices on the surface and a centrally located β-sheet. Both enzymes are traversed by a DNA-binding groove connecting to a uracil-binding pocket, which in the case of MUG is less tailor-made for uracil. Two active site motifs are conserved. In human UNG1 these are 143-GQDPY-147 and 268-HPSPLS-273. These are identical in human, E. coli, yeast and herpes viral proteins. The corresponding motifs in MUG are 16-GINPGL-20, which is identical to the corresponding motif in TDG, and 140-NPSGLS-145, which is related to the corresponding motif in human TDG (MPSSSS). While aspartate in the first motif of UNG has a function in activating water for attack on the C1′ of deoxyribose, asparagine in MUG/TDG can not activate the nucleophilic water, although it may bind and position it. The two conserved motifs in UNG and MUG/TDG have very similar conformations. The active sites of UNG and MUG/TDG are thus related, but specific differences also explain the broader substrate specificity of MUG/TDG. In both cases, the base can only be accommodated in the catalytic pocket after the nucleotide is flipped out of the helix. In UNG, binding of T is sterically hindered by the bulky side chain of Tyr147, which would clash with the 5′-methyl group of T. In the corresponding position in MUG, the amino acid is Gly20 which does not represent a major barrier to thymine. The preference for U : G over T : G is probably explained by position of the side chain in Ser23, the OH-group of which would clash with the 5′ methyl-group of thymine. However, the Ser side chain is free to rotate. Thus, unlike UNG where the Tyr side chain is fixed, there is no rigid structure in MUG preventing binding of pyrimidines with substitutions in the 5′ position. In mammalian TDG, Ala is in the position corresponding to Ser23 in MUG. The smaller side chain of Ala represents an even smaller barrier to binding of T. This probably explains why MUG has a stronger preference for U : G over T : G than TDG (Barrett et al., 1999). Interestingly, substitution of Tyr147 in human UNG with Ala, Ser or Cys results in enzymes that release T and U, but not C, from DNA. Although the catalytic turnover is reduced compared with wild type UNG, it is still higher than for TDG (Kavli et al., 1996).

The discrimination between cytosine and thymine/uracil is less obvious. Possibly, the hydrophobic nature of the catalytic pocket, the specificity for G in the complementary strand, and a weak push that can only flip out unstable base pairs may explain the selectivity for U/T : G over the more stable C : G. The hydrophobic εC would be even better accommodated in the pocket, and it is weakly bonded to the complementary G, explaining why it is a good substrate for TDG (Barrett et al., 1999).

Like several other DNA glycosylases, TDG has N-terminal and C-terminal extensions not present in the bacterial homologues. These extensions may be involved in subcellular sorting and protein–protein interactions, although this has not yet been demonstrated. Yeast two-hybrid screening surprisingly revealed interaction between TDG and the retinoid acid receptor (RAR), as well as retinoid X receptor (RXR). RAR and RXR are intracellular receptors that after binding of ligand function as transcription factors. In fact, in vitro studies have demonstrated that TDG stimulated RAR/RXR-dependent transactivation (Um et al., 1998). Furthermore, TDG has been shown to be a strong repressor of thyroid transcription factor-1 (TTF-1) transcriptional activity (Missero et al., 2001), indicating that TDG may function both as a positive and negative regulator of transcription. Whether the transcription-associated activity of TDG is modulated by sumoylation is presently not known. This, however merits further investigation, since several transcription factors appear to be directly or indirectly regulated by sumoylation (reviewed by Muller et al., 2001).

In conclusion, TDG removes 5-FU, εC, U, T and 5-hmeU in this order from double stranded DNA, preferentially when mismatched to G. In addition, it may function as a transcription factor. Which of these functions is the most important remains unclear.

MBD4 (MED1) – A protein containing a methyl binding domain and a separate glycosylase domain

MBD4 (also called MED1) has central properties in common with TDG, it binds to T : G or U : G mismatches preferentially in CpG contexts and releases the pyrimidine base with low efficiency. Like TDG, it also removes 5-FU and, weakly, εC (reviewed in Bellacosa, 2001). TDG and MBD4 therefore apparently have very similar substrate specificities. Possibly MBD4 through its specific methyl-binding domain may have a more important role in methylated CpG contexts than TDG. MBD4 may also have an additional role in mismatch repair through its interaction with MLH1 (reviewed in Bellacosa, 2001). Furthermore, there is significant evidence associating defects in the MBD4 gene with human cancer (reviewed in Hardeland et al., 2001) while similar evidence is lacking for TDG. The gene for MBD4 has been mapped to position 3q21.22 (Bellacosa, 2001). It is thus the only one of the four uracil-removing proteins that is not located to human chromosome 12 and it is apparently unrelated to the other uracil-DNA glycosylases.

Human MBD4 was discovered as one member of a family of proteins that bind specifically to methylated DNA in vitro (Hendrich and Bird, 1998). MBD4 was also found to interact with the human mismatch repair protein MLH1 in a yeast two-hybrid system. It was also found to have a domain with homology to glycosylases/lyases and displayed an apparent endonuclease activity (Bellacosa et al., 1999). However, MBD4 was later shown to be a monofunctional G : T and U : G mismatch glycosylase (Hendrich et al., 1999). The 580 amino acid protein contains an N-terminal methyl-binding domain, MBD (residues 82–147) and a C-terminal glycosylase domain (residues 401–580) with homology to E. coli thymine glycol glycosylase (EndoIII) and 8-oxoG : A specific adenine glycosylase (MutY). A CpG sequence context was preferred, but not absolutely required since mismatches in other sequences were processed at lower rates (Hendrich et al., 1999). Furthermore, the methylation status of CpGTpG mismatches or CpGUpG mismatches did not affect the glycosylase activity of MBD4 under certain reaction conditions. Nevertheless, the enzyme binds more strongly to 5-meCpGTpG mismatches than to unmethylated mismatches or 5-meCpGmeCpG matches. It was therefore proposed that the function of MBD4 is to counteract the mutagenic effects of deamination of 5-meC to thymine. In this function, the MBD domain may be responsible for the binding specificity and the glycosylase domain for catalysis (Hendrich and Bird, 2000).

Surprisingly, MBD4 has also been reported to have 5-meC DNA glycosylase activity, especially in hemimethylated DNA. It might therefore contribute to active changing of the methylation status in cells (Zhu et al., 2000a). This, as well as the mismatch repair function, are potential functions that could be relevant to the possible role of MBD4 in cancer prevention (reviewed by Bellacosa, 2001). Furthermore, MBD4 mutations were reported in 14 cases of human cancer with microsatellite instability, but not in tumour without microsatellite instability. All of the mutations were deletions of A (one or two) in a polyA track, resulting in frame shift and generation of a stop codon. While six of the cancers also carried a mutation in MLH1, with which MBD4 interacts, five of the tumours did not carry MLH1 mutations and in three the MLH1/MSH2 status was not known (Riccio et al., 1999). These findings suggest that MBD4 may be important in prevention of cancer.

A fifth family of uracil DNA-glycosylases

Recently, a new uracil-DNA glycosylase was identified in the crenarchaeon Pyrobaculum aerophilum by using a search algorithm targeted to the catalytic motifs I and III (Aravind and Koonin, 2000) of the UDG superfamily (Sartori et al., 2002). This UDG, called Pa-UDGb, surprisingly contained no polar residue in the catalytic motif I (GLAPA) which is proposed to activate or orient water in the other UDGs. The enzyme furthermore had a unusual broad substrate specificity, including U, 5-HmU, 5-FU and εC. In addition, Pa-UDGb was able to remove hypoxanthine, and thus is the first member of the UDG superfamily able to remove both pyrimidines and purines (Sartori et al., 2002). The authors thus proposed that Pa-UDGb represented a fifth UDG family, that evolved in organisms living at elevated temperatures to counteract the mutagenic threat of both cytosine and adenine deamination.

Evolution of uracil-DNA glycosylases

The classification of DNA glycosylases into superfamilies, can be based on characteristic sequence motifs as defined in the Pfam database (Bateman et al., 2002). The two largest superfamilies are uracil DNA glycosylases (UDG) and helix–hairpin–helix–GPD glycosylases (HhH–GPD). The HhH–GPD superfamily is named after its hallmark motif (Thayer et al., 1995), a helix–hairpin–helix (HhH) and Gly/Pro rich loop (GP) followed by a conserved aspartate (D). UNG, SMUG1 and TDG belong to the UDG superfamily, which is the most important in terms of uracil processing. MBD4 belongs to the HhH–GPD family, members of which remove a large variety of lesions, including uracil, oxidised bases and certain mismatches, particularly A mismatched to G or 8-oxoG.

The phylogenetic distribution of DNA repair genes has been discussed in detail by Eisen and Hanawalt (1999), while Aravind and Koonin (2000) have specifically analysed the UDG superfamily. The HhH–GPD glycosylases are widespread in both Archaea, bacteria and eukaryotes, and are believed to represent a very ancient gene family. However, the UDG superfamily shows a more non-even distribution pattern. TDG is the most widespread UDG gene family, as it is found both in Archaea, bacteria and eukaryotes. It is distantly related to the AUDG gene family, which is mainly found in Archaea and bacteria, and it seems likely that they share an ancient ancestor. Neither the UNG nor the SMUG1 gene families are found in Archaea. SMUG1 has so far been found only in some eukaryotes. Based on the conservation pattern in the minor-groove intercalation loop it was suggested that SMUG1 may have evolved from an UNG-like enzyme by rapid divergence, possibly to meet special requirements for repair in multicellular animals (Aravind and Koonin, 2000). However, a more traditional phylogenetic analysis tends to place SMUG1 closer to TDG/MUG and AUDG, and more distant from UNG (Sartori et al., 2001). These studies are based on relatively few sequences with low similarity. This makes alignment and analysis difficult, and the interpretation of these data should be done with caution. The origin of SMUG1 therefore remains an open question.

UNG is very widespread in bacteria, and has also been identified in most eukaryotes. It has been suggested that UNG was introduced into eukaryotes by horizontal gene transfer (Aravind and Koonin, 2000), possibly from the mitochondrial genome (Eisen and Hanawalt, 1999). UNG is also the only of these gene families that is found in a large number of viruses, indicating another possible mechanism for horizontal gene transfer. Furthermore, there are more, and more recently established pseudogenes from UNG than from other DNA glycosylases. A Blast search (Altschul et al., 1990) of the Ensembl human genome sequence (Hubbard et al., 2002) identifies at least four certain UNG pseudogenes, compared to 0–2 pseudogenes for other human DNA glycosylase genes (F Drabløs, unpublished).

Sequence comparison of the different gene families within the UDG superfamily identifies several conserved sequence motifs, indicating a common 3D-fold for all UDG-type proteins. This is confirmed by the known 3D structures for UNG and TDG/MUG, as they share a common 3D-fold classified as an alpha beta 3-layer (aba) sandwich in the CATH classification system (Pearl et al., 2000). A direct structural superposition using the DALI software (Holm and Sander, 1993) for MUG (1mug) vs UNG (3eug) gives a highly significant probability score (Z-score 7.3) with a residual means score (RMS) or the 3D structures of 4.0 Å over 145 residues, despite just 13% sequence identity in the corresponding sequence alignment. It therefore seems realistic to assume that all UDG-type proteins share this common fold.

There are at least five relevant sequence motifs in the UDG sequences that can be related to structural features in UNG (Parikh et al., 2000a). The first is the water-activating motif (GQDPYH in human UNG). It is also identified as Motif 1 by Pearl (2000) or Motif I by Aravind and Koonin (2000), and is relatively well conserved in all UDG sequences. This loop coordinates the catalytic-centre water molecule just below the C1′ deoxyribose. The proline-rich motif (PPPPSL) which is directly involved in DNA interaction in UNG is not well-conserved in other UDG sequences. However, when comparing the UNG and MUG structures the main difference is an extra loop outside the actual binding region in UNG. The essential part of the proline-rich motif that is in direct contact with the DNA backbone is structurally conserved in MUG. The uracil-recognition motif (GVLLLN) is conserved mainly in the UNG proteins. However, the actual region seems to be well conserved in most sequences, the variation is mainly with respect to the specific residues found at each position. The motif contributes to uracil recognition by hydrogen bonding to polar atoms of the uracil ring. In the uracil binding pocket there is also a favourable stacking interaction between uracil and a well-conserved phenylalanine residue found between the water-activating and the proline-rich motifs, in addition to the tyrosine from the water-activating motif mentioned above. The uracil-recognition motif is then followed by the glycine-serine motif (LWGS), which is identified as Motif II by Aravind and Koonin. This motif is also involved in DNA interaction. In particular the glycine is well conserved, probably because a side chain at this position would interfere with the close contact between the protein and the DNA backbone. The final motif is the minor-groove intercalation loop motif (HPSPLS), identified as Motif 2 by Pearl and Motif III by Aravind and Koonin. This motif shows some variation, but the histidine and the first proline are conserved in most sequences, except in SMUG1. The histidine is one of the active site residues, and forms hydrogen bond with uracil.

The crystal structures show that UNG has a highly conserved, single domain α/β topology with a central, four-stranded parallel β-sheet surrounded by eight α-helices, classified as an alpha beta 3-layer (aba) sandwich in CATH. This architecture family is a very large one, with 70 topologies listed in CATH. The central β-sheet consists of parallel strands in the order 2-1-3-4.

The HhH-GPD superfamily, to which MBD4 belongs, has a very different architecture. It consists of two α-helical domains classified as mainly alpha orthogonal bundles by CATH. Some members of the HhH–GPD superfamily, including MBD4, also contain a methyl-CpG binding domain (MBD) which has an alpha beta 2-layer sandwich architecture. It has been shown that MBD4 acts as a G : T and G : U specific thymine and uracil glycosylase (Hendrich et al., 1999; Petronzelli et al., 2000). The 3D structure for MBD4 is not known. However, structures are available for several other members of this superfamily, and it can be assumed that important structural features are conserved. The structure of human OGG1 in complex with oxoGC-containing DNA (1ebm) (Bruner et al., 2000) and of Methanobacterium thermoformicicum MIG (1kea) (Mol et al., 2002) can be used as examples. The HhH–GPD-fold consists of a four-helix bundle domain and a six-helix barrel domain, with the active site and the HhH motif located at the interface between these domains. Whereas most DNA-binding proteins seem to use a charged surface rich in lysine and arginine residues to bind backbone phosphates, the DNA binding surface of OGG1 is nearly charge neutral. However, a large number of α-helices have the N-terminal end oriented towards the DNA, maximizing helix-dipole interactions. Only one helix, which is part of the helix–hairpin–helix (HhH) motif, is actually in contact with DNA. As seen in other DNA glycosylases the DNA residue is fully extruded from the DNA helix and inserted into the active-site pocket of the enzyme.

The functional roles of the different glycosylases and coupling to downstream steps in BER

UNG1 and UNG2, together with SMUG1 are the only known DNA glycosylases with preference for single-stranded DNA. UNG-proteins are highly selective for uracil, but remove 5-fluorouracil and certain oxidised pyrimidines with very low efficiency (Krokan et al., 1997). While U in U : G mispairs is generally preferred over U : A, the preference is dependent on the sequence context (Eftedal et al., 1993). The efficient removal of uracil from single-stranded DNA is puzzling since it leaves a non-informative lesion without the information in a complementary strand. Single-stranded DNA is probably mainly found temporarily in transcribed genes and very close to the replication fork. Abasic sites resulting from uracil-removal in single-stranded DNA at the replication fork could be handled by at least three different mechanisms; (i) regression of the replication fork and repair by short patch or long patch BER, (ii) recombination repair using the old strand at the other side of the fork, (iii) translesion DNA synthesis. Regression of a replication fork stalled at a single-strand lesion is well established in E. coli and is an ATP-requiring process. It may in principle apply to all types of lesions that stall the replication fork, including abasic sites (Robu et al., 2001). Recombination using information from the sister chromatid at stalled replication forks (Gruss and Michel, 2001), as well as translesion synthesis across abasic sites are well established processes in bacteria. Interestingly, repair of abasic sites in chromosomal DNA in E. coli in vivo has been demonstrated to require both BER, recombination repair and translesion synthesis (Otterlei et al., 2000). Therefore, abasic sites resulting from the action of UNG at the replication fork is possibly unlikely to be dealt with by BER alone.

After removal of uracil by a uracil-DNA glycosylase and cleavage of the resulting abasic site by AP-endonuclease (APE1/HAP1), the BER pathway splits into two branches. For more comprehensive overviews, the reader should consult other recent reviews (Dogliotti et al., 2001). The presumed major track is the short patch pathway. It uses the dRPase activity of DNA polymerase β to cleave 3′ of the abasic site, thus releasing deoxyribose-5-phosphate. Then Pol β inserts C or T, depending on the template base. Finally DNA ligase 3 seals the nick, aided by the scaffold protein XRCC1. The alternative long patch pathway largely uses replication proteins (Dogliotti et al., 2001) and may take place in replication foci (Otterlei et al., 1999). This pathway requires Pol ε and/or δ, as well as the trimeric sliding clamp and polymerase processivity factor proliferating cell nuclear antigen (PCNA) and the clamp loader replication factor C (RFC). Repair synthesis is stimulated by Pol β, which may be important in the first step of polymerization. The structure specific endonuclease FEN1 removes the 2–8 nucleotide displaced ‘flap’ of DNA and DNA ligase 1 seals the nick (Dogliotti et al., 2001).

As shown in Figure 2, uracil in DNA may be present in different positions relative to a replication fork, and in addition the sequence context may vary. It seems likely that the type of uracil-DNA glycosylase, as well as the mechanism of repair in the subsequent steps will depend on these factors. Uracil present in the fork prior to replication, e.g. in the G1-phase, may mainly result from cytosine deamination and may in principle be removed by any of the four identified human uracil-DNA glycosylases. Most likely, TDG and MBD4 mainly function in CpG-contexts, while SMUG1 and UNG2 may operate in any sequence context, albeit with varying efficiency. Thus, UNG2/UNG1 are known to be less active in GC-rich regions as compared with AT-rich regions (Eftedal et al., 1993, 1994; Slupphaug et al., 1995). Yet, even in such sequences, the catalytic efficiencies of UNG2 and SMUG1 are much higher than those of TDG or MBD4, so the latter enzymes presumably have a limited significance in uracil-repair except in special sequence contexts. SMUG1 was recently suggested to be a main enzyme in repair of uracil in U : G mismatches (Nilsen et al., 2001). However, our data strongly suggest that even in repair of U : G mispairs resulting from cytosine deamination, UNG2 may be a major player (Kavli et al., 2002). UNG2 has at least as low Km as SMUG1, and is present in replication foci as well as in the nucleoplasm. However, UNG2 is essentially excluded from nucleoli, whereas SMUG1 accumulates in nucleoli, suggesting that SMUG1 may have a specialized role in uracil-repair in nucleoli (Kavli et al., 2002). Among the uracil-DNA glycosylases, only UNG2 specifically accumulates in the replication foci during S phase, and as discussed, all experimental evidence suggests that UNG2 has an important role in the removal of misincorporated uracil in replication foci.

Figure 2

Uracil in different positions relative to the replication fork, and proposed mechanisms of repair in different positions

One important problem is the following: How are deaminated cytosines that escape repair prior to replication repaired, if at all? If uracil escapes repair and directs incorporation of adenine, subsequent removal of uracil by UNG and replacement by thymine would result in a G : C→A : T transition, and this would be a poor strategy. However, UNG2 efficiently removes uracil from single-stranded DNA and may thus generate an abasic site that blocks replication. The stalled replication fork may recruit proteins required for fork regression and homologous recombination, which are alternative mechanisms to short patch repair and long patch repair in the downstream steps subsequent to uracil removal. Fortunately, APE1 requires double-stranded DNA and thus the danger of creating a double-strand break is small. Involvement of recombination in the repair of abasic sites has been documented for E. coli (Otterlei et al., 2000). Furthermore, induction of deamination of cytosine by NO is strongly cytotoxic in E. coli cells deficient in recombination (Spek et al., 2001). The finding that recombination factors are required for processing of abasic sites in bacteria suggests that this may also be the case in mammalian cells, since this basic process is highly likely to be conserved. We therefore propose that uracil in single-stranded DNA at the replication fork is incised by UNG2 and repaired by recombination or fork regression, which are both processes requiring recombination proteins.


  1. Altschul SF, Gish W, Miller W, Myers EW, Lipman DJ . 1990 J. Mol. Biol. 215: 403–410

  2. Aravind L, Koonin EV . 2000 Genome Biol. 1: RESEARCH0007

  3. Baker D, Liu P, Burdzy A, Sowers LC . 2002 Chem. Res. Toxicol. 15: 33–39

  4. Barrett TE, Savva R, Panayotou G, Barlow T, Brown T, Jiricny J, Pearl LH . 1998 Cell 92: 117–129

  5. Barrett TE, Scharer OD, Savva R, Brown T, Jiricny J, Verdine GL, Pearl LH . 1999 EMBO J. 18: 6599–6609

  6. Bartsch H, Nair J . 2000 Toxicology 153: 105–114

  7. Basu AK, Wood ML, Niedernhofer LJ, Ramos LA, Essigmann JM . 1993 Biochemistry 32: 12793–12801

  8. Bateman A, Birney E, Cerruti L, Durbin R, Etwiller L, Eddy SR, Griffiths-Jones S, Howe KL, Marshall M, Sonnhammer ELL . 2002 Nucleic Acids Res. 30: 276–280

  9. Bellacosa A . 2001 J. Cell Physiol. 187: 137–144

  10. Bellacosa A, Cicchillitti L, Schepis F, Riccio A, Yeung AT, Matsumoto Y, Golemis EA, Genuardi M, Neri G . 1999 Proc. Natl. Acad. Sci. USA 96: 3969–3974

  11. Bellamy SR, Baldwin GS . 2001 Nucleic Acids Res. 29: 3857–3863

  12. Bennett SE, Sanderson RJ, Mosbaugh DW . 1995 Biochemistry 34: 6109–6119

  13. Bharati S, Krokan HE, Kristiansen L, Otterlei M, Slupphaug G . 1998 Nucleic Acids Res. 26: 4953–4959

  14. Boorstein RJ, Cummings Jr A, Marenstein DR, Chan MK, Ma Y, Neubert TA, Brown SM, Teebor GW . 2001 J. Biol. Chem. 276: 41991–41997

  15. Boorstein RJ, Levy DD, Teebor GW . 1987 Mutat. Res. 183: 257–263

  16. Bouhamdan M, Benichou S, Rey F, Navarro JM, Agostini I, Spire B, Camonis J, Slupphaug G, Vigne R, Benarous R, Sire J . 1996 J. Virol. 70: 697–704

  17. Brown TC, Jiricny J . 1987 Cell 50: 945–950

  18. Bruner SD, Norman DPG, Verdine GL . 2000 Nature 403: 859–866

  19. Cannon Carlson SV, Gokhale H, Teebor GW . 1989 J. Biol. Chem. 264: 13306–13312

  20. Di Noia J, Neuberger MS . 2002 Nature 419: 43–48

  21. Dinner AR, Blackburn GM, Karplus M . 2001 Nature 413: 752–755

  22. Dogliotti E, Fortini P, Pascucci B, Parlanti E . 2001 Prog. Nucleic Acids Res. Mol. Biol. 68: 3–27

  23. Drohat AC, Xiao G, Tordova M, Jagadeesh J, Pankiewicz KW, Watanabe KA, Gilliland GL, Stivers JT . 1999 Biochemistry 38: 11876–11886

  24. Eftedal I, Guddal PH, Slupphaug G, Volden G, Krokan HE . 1993 Nucleic Acids Res. 21: 2095–2101

  25. Eftedal I, Volden G, Krokan HE . 1994 Ann. NY Acad. Sci. 726: 312–314

  26. Eisen JA, Hanawalt PC . 1999 Mutat. Res. 435: 171–213

  27. Faure H, Mousseau M, Cadet J, Guimier C, Tripier M, Hida H, Favier A . 1998 Free Radical Res. 28: 377–382

  28. Friedberg EC, Walker GC, Siede W . 1995 DNA Repair and Mutagenesis Washington, DC: ASM Press pp. 698

  29. Gruss A, Michel B . 2001 Curr. Opin. Microbiol. 4: 595–601

  30. Handa P, Roy S, Varshney U . 2001 J. Biol. Chem. 276: 17324–17331

  31. Hang B, Medina M, Fraenkel-Conrat H, Singer B . 1998 Proc. Natl. Acad. Sci. USA 95: 13561–13566

  32. Haracska L, Johnson RE, Unk I, Phillips BB, Hurwitz J, Prakash L, Prakash S . 2001 Proc. Natl. Acad. Sci. USA 98: 14256–14261

  33. Hardeland U, Bentele M, Jiricny J, Schar P . 2000 J. Biol. Chem. 275: 33449–33456

  34. Hardeland U, Bentele M, Lettieri T, Steinacher R, Jiricny J, Schar P . 2001 Prog. Nucleic Acids Res. Mol. Biol. 68: 235–253

  35. Hardeland U, Steinacher R, Jiricny J, Schar P . 2002 EMBO J. 21: 1456–1464

  36. Haug T, Skorpen F, Aas PA, Malm V, Skjelbred C, Krokan HE . 1998 Nucleic Acids Res. 26: 1449–1457

  37. Haushalter KA, Todd Stukenberg MW, Kirschner MW, Verdine GL . 1999 Curr. Biol. 9: 174–185

  38. Hendrich B, Bird A . 1998 Mol. Cell. Biol. 18: 6538–6547

  39. Hendrich B, Bird A . 2000 DNA Methylation and Cancer 249: 55–74

  40. Hendrich B, Hardeland U, Ng HH, Jiricny J, Bird A . 1999 Nature 401: 301–304

  41. Holm L, Sander C . 1993 J. Mol. Biol. 233: 123–138

  42. Horsfall MJ, Borden A, Lawrence CW . 1997 J. Bact. 179: 2835–2839

  43. Hubbard T, Barker D, Birney E, Cameron G, Chen Y, Clark L, Cox T, Cuff J, Curwen V, Down T, Durbin R, Eyras E, Gilbert J, Hammond M, Huminiecki L, Kasprzyk A, Lehvaslaiho H, Lijnzaad P, Melsopp C, Mongin E, Pettett R, Pocock M, Potter S, Rust A, Schmidt E, Searle S, Slater G, Smith J, Spooner W, Stabenau A, Stalker J, Stupka E, Ureta-Vidal A, Vastrik I, Clamp M . 2002 Nucleic Acids Res. 30: 38–41

  44. Jiang YL, Stivers JT . 2001 Biochemistry 40: 7710–7719

  45. Kavli B, Slupphaug G, Mol CD, Arvai AS, Petersen SB, Tainer JA, Krokan HE . 1996 EMBO J. 15: 3442–3447

  46. Kavli B, Sundheim O, Akbari M, Otterlei M, Nilsen H, Skorpen F, Aas PA, Hagen L, Krokan HE, Slupphaug G . 2002 J. Biol. Chem. (In press). Electronic publication ahead of print, August 2, 2002

  47. Krokan HE, Otterlei M, Nilsen H, Kavli B, Skorpen F, Andersen S, Skjelbred C, Akbari M, Aas PA, Slupphaug G . 2001 Prog. Nucleic Acids Res. Mol. Biol. 68: 365–386

  48. Krokan HE, Standal R, Slupphaug G . 1997 Biochem. J. 325: 1–16

  49. Levy DD, Teebor GW . 1991 Nucleic Acids Res. 19: 3337–3343

  50. Lindahl T . 1974 Proc. Natl. Acad. Sci. USA 71: 3649–3653

  51. Lindahl T . 1993 Nature 362: 709–715

  52. Mellac S, Fazakerley GV, Sowers LC . 1993 Biochemistry 32: 7779–7786

  53. Missero C, Pirro MT, Simeone S, Pischetola M, Di Lauro R . 2001 J. Biol. Chem. 276: 33569–33575

  54. Mol CD, Arvai AS, Begley TJ, Cunningham RP, Tainer JA . 2002 J. Mol. Biol. 315: 373–384

  55. Mol CD, Arvai AS, Slupphaug G, Kavli B, Alseth I, Krokan HE, Tainer JA . 1995 Cell 80: 869–878

  56. Moriya M, Zhang W, Johnson F, Grollman AP . 1994 Proc. Natl. Acad. Sci. USA 91: 11899–11903

  57. Muller S, Hoege C, Pyrowolakis G, Jentsch S . 2001 Nature Rev. Mol. Cell Biol. 2: 202–210

  58. Muramatsu M, Kinoshita K, Fagarasan S, Yamada S, Shinkai Y, Honjo T . 2000 Cell 102: 553–563

  59. Muramatsu M, Sankaranand VS, Anant S, Sugai M, Kinoshita K, Davidson NO, Honjo T . 1999 J. Biol. Chem. 274: 18470–18476

  60. Neddermann P, Gallinari P, Lettieri T, Schmid D, Truong O, Hsuan JJ, Wiebauer K, Jiricny J . 1996 J. Biol. Chem. 271: 12767–12774

  61. Neddermann P, Jiricny J . 1993 J. Biol. Chem. 268: 21218–21224

  62. Nilsen H, Haushalter KA, Robins P, Barnes DE, Verdine GL, Lindahl T . 2001 EMBO J. 20: 4278–4286

  63. Nilsen H, Otterlei M, Haug T, Solum K, Nagelhus TA, Skorpen F, Krokan HE . 1997 Nucleic Acids Res. 25: 750–755

  64. Nilsen H, Rosewell I, Robins P, Skjelbred CF, Andersen S, Slupphaug G, Daly G, Krokan HE, Lindahl T, Barnes DE . 2000 Mol. Cell 5: 1059–1065

  65. Nilsen H, Yazdankhah SP, Eftedal I, Krokan HE . 1995 FEBS Lett. 362: 205–209

  66. Olsen LC, Aasland R, Wittwer CU, Krokan HE, Helland DE . 1989 EMBO J. 8: 3121–3125

  67. Otterlei M, Haug T, Nagelhus TA, Slupphaug G, Lindmo T, Krokan HE . 1998 Nucleic Acids Res. 26: 4611–4617

  68. Otterlei M, Kavli B, Standal R, Skjelbred C, Bharati S, Krokan HE . 2000 EMBO J. 19: 5542–5551

  69. Otterlei M, Warbrick E, Nagelhus TA, Haug T, Slupphaug G, Akbari M, Aas PA, Steinsbekk K, Bakke O, Krokan HE . 1999 EMBO J. 18: 3834–3844

  70. Parikh SS, Mol CD, Slupphaug G, Bharati S, Krokan HE, Tainer JA . 1998 EMBO J. 17: 5214–5226

  71. Parikh SS, Putnam CD, Tainer JA . 2000a Mutat. Res. 460: 183–199

  72. Parikh SS, Walcher G, Jones GD, Slupphaug G, Krokan HE, Blackburn GM, Tainer JA . 2000b Proc. Natl. Acad. Sci. USA 97: 5083–5088

  73. Pasternack LB, Bramham J, Mayol L, Galeone A, Jia X, Kearns DR . 1996 Nucleic Acids Res. 24: 2740–2745

  74. Pearl FMG, Lee D, Bray JE, Sillitoe I, Todd AE, Harrison AP, Thornton JM, Orengo CA . 2000 Nucleic Acids Res. 28: 277–282

  75. Pearl LH . 2000 Mutat. Res. 460: 165–181

  76. Petersen-Mahrt SK, Harris RS, Neuberger MS . 2002 Nature 418: 99–103

  77. Petronzelli F, Riccio A, Markham GD, Seeholzer SH, Stoerker J, Genuardi M, Yeung AT, Matsumoto Y, Bellacosa A . 2000 J. Biol. Chem. 275: 32422–32429

  78. Radany EH, Dornfeld KJ, Sanderson RJ, Savage MK, Majumdar A, Seidman MM, Mosbaugh DW . 2000 Mutat. Res. 461: 41–58

  79. Riccio A, Aaltonen LA, Godwin AK, Loukola A, Percesepe A, Salovaara R, Masciullo V, Genuardi M, Paravatou-Petsotas M, Bassi DE, Ruggeri BA, Klein-Szanto AJP, Testa JR, Neri G, Bellacosa A . 1999 Nature Genetics 23: 266–268

  80. Robu ME, Inman RB, Cox MM . 2001 Proc. Natl. Acad. Sci. USA 98: 8211–8218

  81. Rossi R, Villa A, Negri C, Scovassi I, Ciarrocchi G, Biamonti G, Montecucco A . 1999 EMBO J. 18: 5745–5754

  82. Rusmintratip V, Sowers LC . 2000 Proc. Natl. Acad. Sci. USA 97: 14183–14187

  83. Saparbaev M, Laval J . 1998 Proc. Natl. Acad. Sci. USA 95: 8508–8513

  84. Sartori AA, Fitz-Gibbon S, Hanjing Y, Miller JH, Jiricny J . 2002 EMBO J. 21: 3182–3191

  85. Sartori AA, Schar P, Fitz-Gibbon S, Miller JH, Jiricny J . 2001 J. Biol. Chem. 276: 29979–29986

  86. Savva R, McAuley Hecht K, Brown T, Pearl L . 1995 Nature 373: 487–493

  87. Sekiguchi M, Hayakawa M, Makino F, Tanaka K, Okada Y . 1976 Biochem. Biophys. Res. Commun. 73: 293–299

  88. Shapiro R . 1980 Chromosome damage and repair Seeberg E, Kleppe K (ed) New York: Plenum Press pp. 3–18

  89. Sheehy AM, Gaddis NC, Choi JD, Malim MH . 2002 Nature 418: 646–650

  90. Shroyer MJ, Bennett SE, Putnam CD, Tainer JA, Mosbaugh DW . 1999 Biochemistry 38: 4834–4845

  91. Slupphaug G, Eftedal I, Kavli B, Bharati S, Helle NM, Haug T, Levine DW, Krokan HE . 1995 Biochemistry 34: 128–138

  92. Slupphaug G, Mol CD, Kavli B, Arvai AS, Krokan HE, Tainer JA . 1996 Nature 384: 87–92

  93. Spek EJ, Wright TL, Stitt MS, Taghizadeh NR, Tannenbaum SR, Marinus MG, Engelward BP . 2001 J. Bact. 183: 131–138

  94. Stivers JT, Pankiewicz KW, Watanabe KA . 1999 Biochemistry 38: 952–963

  95. Thayer MM, Ahern H, Xing D, Cunningham RP, Tainer JA . 1995 EMBO J. 14: 4108–4120

  96. Tu YQ, Dammann R, Pfeifer GP . 1998 J. Mol. Biol. 284: 297–311

  97. Tye BK, Nyman PO, Lehman IR, Hochhauser S, Weiss B . 1977 Proc. Natl. Acad. Sci. USA 74: 154–157

  98. Um S, Harbers M, Benecke A, Pieratt B, Losson R, Chambon P . 1998 J. Biol. Chem. 273: 20728–20736

  99. Vidal AE, Hickson ID, Boiteux S, Radicella JP . 2001 Nucleic Acids Res. 29: 1285–1292

  100. Vilpo JA, Vilpo LM . 1995 Mutat. Res. 316: 123–131

  101. Waters TR, Gallinari P, Jiricny J, Swann PF . 1999 J. Biol. Chem. 274: 67–74

  102. Waters TR, Swann PF . 2000 Mutat. Res. 462: 137–147

  103. Werner RM, Jiang YL, Gordley RG, Jagadeesh GJ, Ladner JE, Xiao G, Tordova M, Gilliland GL, Stivers JT . 2000 Biochemistry 39: 12585–12594

  104. Werner RM, Stivers JT . 2000 Biochemistry 39: 14054–14064

  105. Willetts KE, Rey F, Agostini I, Navarro JM, Baudat Y, Vigne R, Sire J . 1999 J. Virol. 73: 1682–1688

  106. Wist E, Unhjem O, Krokan HE . 1978 Biochim. Biophys. Acta 250: 253–270

  107. Wittwer CU, Bauw G, Krokan HE . 1989 Biochemistry 28: 780–784

  108. Wong I, Lundquist AJ, Bernards AS, Mosbaugh DW . 2002 J. Biol. Chem. 277: 19424–19432

  109. Xiao G, Tordova M, Jagadeesh J, Drohat AC, Stivers JT, Gilliland GL . 1999 Proteins 35: 13–24

  110. Zhu B, Zheng Y, Angliker H, Schwarz S, Thiry S, Siegmann M, Jost JP . 2000a Nucleic Acids Res. 28: 4157–4165

  111. Zhu B, Zheng Y, Hess D, Angliker H, Schwarz S, Siegmann M, Thiry S, Jost JP . 2000b Proc. Natl. Acad. Sci. USA 97: 5135–5139

Download references


This work was sponsored by The Norwegian Cancer Society, The Research Council of Norway, The Cancer Fund at the St. Olavs Hospital, Trondheim and The Svanhild and Arne Must Fund for Medical Research.

Author information

Correspondence to Hans E Krokan.

Rights and permissions

Reprints and Permissions

About this article

Cite this article

Krokan, H., Drabløs, F. & Slupphaug, G. Uracil in DNA – occurrence, consequences and repair. Oncogene 21, 8935–8948 (2002). https://doi.org/10.1038/sj.onc.1205996

Download citation


  • uracil
  • uracil-DNA glycosylase
  • DNA repair

Further reading