Advertisement

Eukaryotic Lagging Strand DNA Replication Employs a Multi-pathway Mechanism That Protects Genome Integrity*

  • Lata Balakrishnan
    Affiliations
    Department of Biochemistry and Biophysics, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642
    Search for articles by this author
  • Robert A. Bambara
    Correspondence
    To whom correspondence should be addressed
    Affiliations
    Department of Biochemistry and Biophysics, University of Rochester School of Medicine and Dentistry, Rochester, New York 14642
    Search for articles by this author
  • Author Footnotes
    * This work was supported, in whole or in part, by National Institutes of Health Grant GM024441 (to R. A. B.). This minireview will be reprinted in the 2011 Minireview Compendium, which will be available in January, 2012.
Open AccessPublished:December 21, 2010DOI:https://doi.org/10.1074/jbc.R110.209502
      In eukaryotic nuclear DNA replication, one strand of DNA is synthesized continuously, but the other is made as Okazaki fragments that are later joined. Discontinuous synthesis is inherently more complex, and fragmented intermediates create risks for disruptions of genome integrity. Genetic analyses and biochemical reconstitutions indicate that several parallel pathways evolved to ensure that the fragments are made and joined with integrity. An RNA primer is removed from each fragment before joining by a process involving polymerase-dependent displacement into a single-stranded flap. Evidence in vitro suggests that, with most fragments, short flaps are displaced and efficiently cleaved. Some flaps can become long, but these are also removed to allow joining. Rarely, a flap can form structure, necessitating displacement of the entire fragment. There is now evidence that post-translational protein modification regulates the flow through the pathways to favor protection of genomic information in regions of actively transcribed chromatin.

      Introduction

      During the S phase of the cell cycle, the genetic material within the eukaryotic cell is duplicated with high efficiency and accuracy. One might expect nuclear DNA replication, an ancient and fundamental cellular requirement, to be a relatively simple process. In reality, it is very complex, with the evolution of several parallel pathways to ensure the maintenance of genome stability. Based on the 5′–3′-directionality of DNA polymerases, replication proceeds by continuous synthesis on the leading strand, growing in the same direction as the opening of the parental strands, and discontinuous synthesis on the lagging strand, growing in the opposite direction (
      • Kornberg A.
      • Baker T.A.
      ). The lagging strand is first made as short fragments of DNA, which are initiated by RNA/DNA primers. These fragments, ∼100–150 nucleotides (nt)
      The abbreviations used are: nt, nucleotides; pol, DNA polymerase; ssDNA, single-stranded DNA; PCNA, proliferating cell nuclear antigen; RPA, replication protein A; LigI, DNA ligase I; BER, base excision repair; LP-BER, long patch BER; dRP, 5′-deoxyribose phosphate.
      in length, are known as Okazaki fragments (
      • Okazaki R.
      • Okazaki T.
      • Sakabe K.
      • Sugimoto K.
      • Sugino A.
      ,
      • Sakabe K.
      • Okazaki R.
      ). Prokaryotic DNA replication occurs within a fork structure in which the lagging strand loops around in a proposed “trombone model” to help orient the replication enzymes for repeated synthesis and joining (
      • Alberts B.M.
      • Barry J.
      • Bedinger P.
      • Formosa T.
      • Jongeneel C.V.
      • Kreuzer K.N.
      ). A similar structure may be employed by eukaryotes. Synthesis of DNA strands occurs at identical rates, suggesting a coordination between leading and lagging strand synthesis (
      • Pandey M.
      • Syed S.
      • Donmez I.
      • Patel G.
      • Ha T.
      • Patel S.S.
      ).

      Prokaryotic DNA Polymerases

      Both strands of the parental prokaryotic genome are replicated by DNA polymerase (pol) III holoenzyme (reviewed in Ref.
      • Johnson A.
      • O'Donnell M.
      ). A multisubunit complex of 10 components, the α-, ϵ-, and θ-subunits make up the catalytic core, acting to polymerize and edit the newly synthesized DNA. Association with the β-subunit, which encircles the DNA as a sliding clamp, completes the holoenzyme, which carries out highly processive synthesis. The triple-DNA polymerase replisome model suggests that three core polymerases assemble at the replication fork, with one working to extend the leading strand and two synthesizing on the lagging strand DNA. Because synthesis on the lagging strand is more complex, it is possible that the presence of two polymerases might help to coordinate the replication rates on the leading and lagging strands to avoid uncoupling of the replication process (
      • McInerney P.
      • Johnson A.
      • Katz F.
      • O'Donnell M.
      ).
      On the lagging strand, the holoenzyme quickly releases from the template as it encounters the 5′-end of the preceding downstream Okazaki fragment. This has been termed the “collision model.” New lines of evidence from O'Donnell and co-workers (
      • Georgescu R.E.
      • Kurth I.
      • Yao N.Y.
      • Stewart J.
      • Yurieva O.
      • O'Donnell M.
      ) argue against the 5′-end primer recognition to trigger collision release by the polymerase holoenzyme. Instead, it has been proposed that the lack of single-stranded DNA (ssDNA) on encountering the preceding Okazaki fragment triggers polymerase release from the β-clamp and the DNA (
      • Georgescu R.E.
      • Kurth I.
      • Yao N.Y.
      • Stewart J.
      • Yurieva O.
      • O'Donnell M.
      ). Dissociation of pol III results in polymerase switching, whereby pol I takes over the maturation of Okazaki fragments. pol I has intrinsic 5′–3′-exonuclease and polymerase activities, so it can mediate both the removal of RNA primers from the 5′-end and the synthesis of nucleotides onto Okazaki fragments. pol I performs nick translation for RNA removal, a combination of strand displacement synthesis and cleavage of resulting short flaps suggested by Setlow et al. (
      • Setlow P.
      • Brutlag D.
      • Kornberg A.
      ), who showed that products of 5′-exonuclease activity were both mono- and oligonucleotides.

      Eukaryotic DNA Polymerases

      Initial studies of eukaryotic DNA replication used SV40 as a model system. Reconstitution of SV40 replication effectively represents cellular processes because the virus makes extensive use of cellular replication proteins. Work with SV40 suggested that the cell initiates replication using pol α/primase (
      • Waga S.
      • Bauer G.
      • Stillman B.
      ). Consisting of four subunits (p180, p70, p58, and p48), pol α is the only eukaryotic polymerase that displays primase activity. The primase catalyzes the synthesis of ∼8–10 nt of RNA; the DNA polymerase then further extends the initiator segment by adding ∼20–20 nt of DNA (
      • Arezi B.
      • Kuchta R.D.
      ). Lacking a proofreading 3′–5′-exonuclease activity, polymerization by pol α is error-prone.
      The studies with SV40 further indicated that pol δ was the main replicative polymerase on both the leading and lagging viral strands, initiating from the primers made by pol α (
      • Tsurimoto T.
      • Melendy T.
      • Stillman B.
      ). However, a number of recent studies in yeast by Kunkel et al. (
      • Pursell Z.F.
      • Isoz I.
      • Lundström E.B.
      • Johansson E.
      • Kunkel T.A.
      ) have definitively identified pol ϵ as the polymerase responsible for synthesis of the cellular leading strand. Consisting of four subunits (Pol2, Dpb2, Dpb3, and Dpb4), pol ϵ is highly processive and is made more processive by proliferating cell nuclear antigen (PCNA), as appropriate for synthesis on the leading strand (
      • Waga S.
      • Stillman B.
      ).
      After initial priming by pol α, pol δ takes over the synthesis on the lagging strand. In Saccharomyces cerevisiae, pol δ is composed of three subunits (Pol3, Pol31, and Pol32); however, in Schizosaccharomyces pombe and mammalian cells, a fourth subunit functions to stabilize the polymerase holoenzyme. pol δ displays both polymerase and 3′–5′-exonuclease activities. Using its proofreading function, pol δ can correct errors made by pol α, thereby protecting genome stability (
      • Pavlov Y.I.
      • Frahm C.
      • Nick McElhinny S.A.
      • Niimi A.
      • Suzuki M.
      • Kunkel T.A.
      ).

      Lagging Strand Replication Machinery Accessory Factors

      Replication protein A (RPA) assembles on the ssDNA immediately after the helicases unwind double-stranded DNA, creating a replication fork. Binding of RPA protects the ssDNA from cellular nucleases and also prevents formation of hairpin structures that might impede the progression of the replication fork (
      • Wold M.S.
      ). Additionally, RPA coordinates the assembly and disassembly of replication-associated proteins on the ssDNA. pol α/primase recognizes RPA-coated ssDNA and initiates primer synthesis on the DNA template (
      • Wold M.S.
      ). Association of the ATP-dependent replication factor C with pol α triggers the switch from replication initiation to replication elongation, wherein pol α is displaced, and PCNA (a functional homolog of the prokaryotic β-clamp) and pol δ are loaded onto the replicating segment of DNA (
      • Tsurimoto T.
      • Melendy T.
      • Stillman B.
      ). PCNA improves the processivity of pol δ and stimulates strand displacement synthesis (
      • Burgers P.M.
      ).

      Involvement of Okazaki Maturation Proteins in Long Patch Base Excision Repair

      A large majority of DNA repair within the cell is processed by the base excision repair (BER) pathway. Depending on the patch length of repair, BER can be divided into short patch BER (correction of a single nucleotide) or long patch BER (LP-BER; correction of a patch 2–12 nt in length) (
      • Matsumoto Y.
      • Kim K.
      ,
      • Piersen C.E.
      • Prasad R.
      • Wilson S.H.
      • Lloyd R.S.
      ,
      • Wilson S.H.
      ). Many of the mechanisms and protein components of Okazaki fragment maturation are also involved in LP-BER. pol β is the main BER polymerase, containing both synthesis and lyase functions. Excision of a damaged base by APE1 (apurinic/apyrimidinic endonuclease 1) creates a 5′-deoxyribose phosphate (dRP) residue, which is cleaved by the lyase activity of pol β, creating a nicked substrate. DNA ligase III then seals the nick, completing short patch BER. When the dRP residue is either oxidized or reduced, the lyase activity of pol β is inhibited. In this event, pol β performs strand displacement synthesis to shift the dRP residue into a 5′-flap, which is subsequently recognized and cleaved by FEN1. LigI seals the nick in LP-BER (Fig. 2) (
      • Wilson S.H.
      ,
      • Memisoglu A.
      • Samson L.
      ,
      • Sung J.S.
      • Demple B.
      ,
      • Wilson 3rd, D.M.
      • Barsky D.
      ). Although Dna2 has been recently implicated in mitochondrial LP-BER, the function of Dna2 in nuclear LP-BER has not been ascertained (
      • Zheng L.
      • Zhou M.
      • Guo Z.
      • Lu H.
      • Qian L.
      • Dai H.
      • Qiu J.
      • Yakubovskaya E.
      • Bogenhagen D.F.
      • Demple B.
      • Shen B.
      ). Studies using plasmid DNA containing a single base lesion have shown that the repair patch size for LP-BER varies from 6 to 12 nt. This patch length is not long enough to stably bind RPA and push the repair process to require the long flap-processing pathway. However, under specific conditions, as described below, the displaced repair patch length might be sufficient to require proteins for long flap processing.

      Regulation of the Pathways for Okazaki Fragment Maturation and LP-BER by Acetylation

      FEN1, the central component of both the short and long flap pathways, has been reported to be acetylated by p300 acetyltransferase (
      • Hasan S.
      • Stucki M.
      • Hassa P.O.
      • Imhof R.
      • Gehrig P.
      • Hunziker P.
      • Hübscher U.
      • Hottiger M.O.
      ,
      • Friedrich-Heineken E.
      • Henneke G.
      • Ferrari E.
      • Hübscher U.
      ). Acetylation decreases its ability to cleave flap substrates by ∼90% (
      • Hasan S.
      • Stucki M.
      • Hassa P.O.
      • Imhof R.
      • Gehrig P.
      • Hunziker P.
      • Hübscher U.
      • Hottiger M.O.
      ,
      • Friedrich-Heineken E.
      • Henneke G.
      • Ferrari E.
      • Hübscher U.
      ). Haploinsufficiency of FEN1, in which the nuclease is present at 50% of the normal level, has deleterious effects in the cell (
      • Kucherlapati M.
      • Yang K.
      • Kuraguchi M.
      • Zhao J.
      • Lia M.
      • Heyer J.
      • Kane M.F.
      • Fan K.
      • Russell R.
      • Brown A.M.
      • Kneitz B.
      • Edelmann W.
      • Kolodner R.D.
      • Lipkin M.
      • Kucherlapati R.
      ,
      • Zheng L.
      • Dai H.
      • Zhou M.
      • Li M.
      • Singh P.
      • Qiu J.
      • Tsark W.
      • Huang Q.
      • Kernstine K.
      • Zhang X.
      • Lin D.
      • Shen B.
      ). Consequently, an intentional down-regulation of nuclease function by acetylation seems undesirable in the context of genome stability. We recently discovered that Dna2, the functional and structural interacting partner of FEN1, is also post-translationally modified by p300 (
      • Balakrishnan L.
      • Stewart J.
      • Polaczek P.
      • Campbell J.L.
      • Bambara R.A.
      ). However, unlike FEN1, the nuclease, helicase, ATPase, and binding activities of Dna2 are all greatly stimulated by acetylation (
      • Balakrishnan L.
      • Stewart J.
      • Polaczek P.
      • Campbell J.L.
      • Bambara R.A.
      ). Notably, we also found that in a small subset of substrates, Dna2 was able to cleave past the base of the flap, potentially eliminating the need for cleavage by FEN1. Apparently, regulation of FEN1 and Dna2 promotes displacement of more nucleotides into each flap before ligation of adjacent Okazaki fragments.
      Why would the cell want to intentionally displace a greater length of flap? Negative effects include more likely formation of secondary hairpin structures or recombination with other complementary sequences. However, a reasonable hypothesis is that regulation by acetylation has evolved because intentional lengthening of the flap would increase the replication and repair patch lengths that are replaced. Since pol α can, on account of its lack of proofreading activity, occasionally synthesize an error-prone initiator primer, if long flaps are displaced, cleaved, and subsequently ligated, it is more likely that mismatches synthesized by pol α will be removed. As a high fidelity polymerase, pol δ would incorporate correct bases into the longer patch. Increased activity of Dna2 would ensure that as the flaps get long, they are efficiently processed to create FEN1 substrates to allow for proper maturation of the Okazaki fragment.
      Significantly, a recent report on the mass spectrometric analysis of three different cell types for protein acetylation identified many replication and repair proteins, including pol δ (on the p12 subunit) and RPA (on the 70-kDa subunit) (
      • Choudhary C.
      • Kumar C.
      • Gnad F.
      • Nielsen M.L.
      • Rehman M.
      • Walther T.C.
      • Olsen J.V.
      • Mann M.
      ). Initial characterization of acetylated replication/repair proteins suggests that the modification regulates the length of the flaps, consistent with our hypothesis. PCNA was previously known to be modified by p300, improving its ability to bind to the polymerases, especially pol δ (
      • Naryzhny S.N.
      • Lee H.
      ). Because PCNA is the processivity factor for pol δ, on acetylation, PCNA is likely to improve the ability of pol δ to displace strands. Strikingly, our preliminary analyses showed that acetylation of pol δ greatly improved its strand displacement functions on Okazaki fragment substrates.
      L. Balakrishnan, B. van Loon, U. Hübscher, and R. A. Bambara, unpublished data.
      Mechanistic effects of acetylation of these proteins are all consistent with intentional replacement of longer patches in DNA replication and repair. Additionally, both BLM and WRN were shown to be acetylated using mass spectrometric analysis. Acetylation of WRN stimulates its 3′–5′-helicase activity (
      • Muftuoglu M.
      • Kusumoto R.
      • Speina E.
      • Beck G.
      • Cheng W.H.
      • Bohr V.A.
      ). The effect of modification on BLM is currently unknown. A summary of intermediates and products expected in the presence of unmodified and acetylated proteins is depicted in Fig. 3.
      Figure thumbnail gr3
      FIGURE 3The long flap pathway promotes genome stability by increasing the fidelity of the replication process. The dark blue lines on the DNA substrate represent the primer nucleotides added by pol α, and the red stars represent mismatches on the downstream Okazaki DNA sequence.
      With respect to repair, if LP-BER makes a longer patch, it is more likely to remove damage on several adjacent nucleotides. Significantly, BER proteins such as DNA glycosylase OGG1 (
      • Bhakat K.K.
      • Mokkapati S.K.
      • Boldogh I.
      • Hazra T.K.
      • Mitra S.
      ), APE1 (
      • Bhakat K.K.
      • Izumi T.
      • Yang S.H.
      • Hazra T.K.
      • Mitra S.
      ), and pol β (
      • Hasan S.
      • El-Andaloussi N.
      • Hardeland U.
      • Hassa P.O.
      • Bürki C.
      • Imhof R.
      • Schär P.
      • Hottiger M.O.
      ) have also been found to be acetylated by p300. Acetylation of pol β selectively abrogates its dRP lyase activity (
      • Hasan S.
      • El-Andaloussi N.
      • Hardeland U.
      • Hassa P.O.
      • Bürki C.
      • Imhof R.
      • Schär P.
      • Hottiger M.O.
      ). Inhibition of lyase activity would drive the repair into the LP-BER pathway, with the formation of longer repair patch lengths. With a shift toward more LP-BER, it is possible that RPA and Dna2 would be involved in processing these flaps, similar to the situation in Okazaki fragment maturation.
      The long patch replacement hypothesis also suggests that the histone acetyltransferase p300 is enriched in regions of active chromatin not only to improve accessibility of the DNA to transcription machinery but also to promote higher fidelity DNA replication and repair to compensate for the destabilizing effects of transcription. Many of the proteins undergoing acetylation are also post-translationally modified by phosphorylation, methylation, and ubiquitination. Distinctively, the modification by acetylation seems to have the uniform effect of increasing the patch length of DNA replication/repair to improve genome stability. Additionally, other forms of modification are likely to regulate protein functions that interact with the effects of acetylation. For example, phosphorylation of FEN1 disrupts binding to PCNA (
      • Henneke G.
      • Koundrioukoff S.
      • Hübscher U.
      ), resulting in defects in Okazaki fragment ligation. Conversely, arginine methylation of FEN1 suppresses phosphorylation of FEN1, thereby increasing binding affinity for PCNA (
      • Guo Z.
      • Zheng L.
      • Xu H.
      • Dai H.
      • Zhou M.
      • Pascua M.R.
      • Chen Q.M.
      • Shen B.
      ). As we begin to understand the cross-effects among different forms of modifications on replication and repair proteins, we may be able to map out a combinatorial modification code that regulates the proteins and the pathways for highest fidelity to slow the progression of cancer, neurodegenerative disorders, and aging.

      Acknowledgments

      We thank Drs. Ulrich Hübscher and Marc Wold and members of the Bambara laboratory for critical reading of the manuscript. We especially thank Christopher Petrides and Athena Kantartzis for assistance with the figures.

      REFERENCES

        • Kornberg A.
        • Baker T.A.
        DNA Replication. 2nd Ed. W. H. Freeman, New York1992
        • Okazaki R.
        • Okazaki T.
        • Sakabe K.
        • Sugimoto K.
        • Sugino A.
        Proc. Natl. Acad. Sci. U.S.A. 1968; 59: 598-605
        • Sakabe K.
        • Okazaki R.
        Biochim. Biophys. Acta. 1966; 129: 651-654
        • Alberts B.M.
        • Barry J.
        • Bedinger P.
        • Formosa T.
        • Jongeneel C.V.
        • Kreuzer K.N.
        Cold Spring Harbor Symp. Quant. Biol. 1983; 47: 655-668
        • Pandey M.
        • Syed S.
        • Donmez I.
        • Patel G.
        • Ha T.
        • Patel S.S.
        Nature. 2009; 462: 940-943
        • Johnson A.
        • O'Donnell M.
        Annu. Rev. Biochem. 2005; 74: 283-315
        • McInerney P.
        • Johnson A.
        • Katz F.
        • O'Donnell M.
        Mol. Cell. 2007; 27: 527-538
        • Georgescu R.E.
        • Kurth I.
        • Yao N.Y.
        • Stewart J.
        • Yurieva O.
        • O'Donnell M.
        EMBO J. 2009; 28: 2981-2991
        • Setlow P.
        • Brutlag D.
        • Kornberg A.
        J. Biol. Chem. 1972; 247: 224-231
        • Waga S.
        • Bauer G.
        • Stillman B.
        J. Biol. Chem. 1994; 269: 10923-10934
        • Arezi B.
        • Kuchta R.D.
        Trends Biochem. Sci. 2000; 25: 572-576
        • Tsurimoto T.
        • Melendy T.
        • Stillman B.
        Nature. 1990; 346: 534-539
        • Pursell Z.F.
        • Isoz I.
        • Lundström E.B.
        • Johansson E.
        • Kunkel T.A.
        Science. 2007; 317: 127-130
        • Waga S.
        • Stillman B.
        Annu. Rev. Biochem. 1998; 67: 721-751
        • Pavlov Y.I.
        • Frahm C.
        • Nick McElhinny S.A.
        • Niimi A.
        • Suzuki M.
        • Kunkel T.A.
        Curr. Biol. 2006; 16: 202-207
        • Wold M.S.
        Annu. Rev. Biochem. 1997; 66: 61-92
        • Burgers P.M.
        J. Biol. Chem. 1991; 266: 22698-22706
        • Bambara R.A.
        • Murante R.S.
        • Henricksen L.A.
        J. Biol. Chem. 1997; 272: 4647-4650
        • Lieber M.R.
        BioEssays. 1997; 19: 233-240
        • Gloor J.W.
        • Balakrishnan L.
        • Bambara R.A.
        J. Biol. Chem. 2010; 285: 34922-34931
        • Burgers P.M.
        J. Biol. Chem. 2009; 284: 4041-4045
        • Li X.
        • Li J.
        • Harrington J.
        • Lieber M.R.
        • Burgers P.M.
        J. Biol. Chem. 1995; 270: 22109-22112
        • Ayyagari R.
        • Gomes X.V.
        • Gordenin D.A.
        • Burgers P.M.
        J. Biol. Chem. 2003; 278: 1618-1625
        • Rossi M.L.
        • Bambara R.A.
        J. Biol. Chem. 2006; 281: 26051-26061
        • Fanning E.
        • Klimovich V.
        • Nager A.R.
        Nucleic Acids Res. 2006; 34: 4126-4137
        • Bae S.H.
        • Bae K.H.
        • Kim J.A.
        • Seo Y.S.
        Nature. 2001; 412: 456-461
        • Bae K.H.
        • Kim H.S.
        • Bae S.H.
        • Kang H.Y.
        • Brill S.
        • Seo Y.S.
        Nucleic Acids Res. 2003; 31: 3006-3015
        • Stewart J.A.
        • Miller A.S.
        • Campbell J.L.
        • Bambara R.A.
        J. Biol. Chem. 2008; 283: 31356-31365
        • Budd M.E.
        • Reis C.C.
        • Smith S.
        • Myung K.
        • Campbell J.L.
        Mol. Cell. Biol. 2006; 26: 2490-2500
        • Rossi M.L.
        • Pike J.E.
        • Wang W.
        • Burgers P.M.
        • Campbell J.L.
        • Bambara R.A.
        J. Biol. Chem. 2008; 283: 27483-27493
        • Henry R.A.
        • Balakrishnan L.
        • Ying-Lin S.T.
        • Campbell J.L.
        • Bambara R.A.
        J. Biol. Chem. 2010; 285: 28496-28505
        • Matsumoto Y.
        • Kim K.
        Science. 1995; 269: 699-702
        • Piersen C.E.
        • Prasad R.
        • Wilson S.H.
        • Lloyd R.S.
        J. Biol. Chem. 1996; 271: 17811-17815
        • Wilson S.H.
        Mutat. Res. 1998; 407: 203-215
        • Memisoglu A.
        • Samson L.
        Mutat. Res. 2000; 451: 39-51
        • Sung J.S.
        • Demple B.
        FEBS J. 2006; 273: 1620-1629
        • Wilson 3rd, D.M.
        • Barsky D.
        Mutat. Res. 2001; 485: 283-307
        • Zheng L.
        • Zhou M.
        • Guo Z.
        • Lu H.
        • Qian L.
        • Dai H.
        • Qiu J.
        • Yakubovskaya E.
        • Bogenhagen D.F.
        • Demple B.
        • Shen B.
        Mol. Cell. 2008; 32: 325-336
        • Hasan S.
        • Stucki M.
        • Hassa P.O.
        • Imhof R.
        • Gehrig P.
        • Hunziker P.
        • Hübscher U.
        • Hottiger M.O.
        Mol. Cell. 2001; 7: 1221-1231
        • Friedrich-Heineken E.
        • Henneke G.
        • Ferrari E.
        • Hübscher U.
        J. Mol. Biol. 2003; 328: 73-84
        • Kucherlapati M.
        • Yang K.
        • Kuraguchi M.
        • Zhao J.
        • Lia M.
        • Heyer J.
        • Kane M.F.
        • Fan K.
        • Russell R.
        • Brown A.M.
        • Kneitz B.
        • Edelmann W.
        • Kolodner R.D.
        • Lipkin M.
        • Kucherlapati R.
        Proc. Natl. Acad. Sci. U.S.A. 2002; 99: 9924-9929
        • Zheng L.
        • Dai H.
        • Zhou M.
        • Li M.
        • Singh P.
        • Qiu J.
        • Tsark W.
        • Huang Q.
        • Kernstine K.
        • Zhang X.
        • Lin D.
        • Shen B.
        Nat. Med. 2007; 13: 812-819
        • Balakrishnan L.
        • Stewart J.
        • Polaczek P.
        • Campbell J.L.
        • Bambara R.A.
        J. Biol. Chem. 2010; 285: 4398-4404
        • Choudhary C.
        • Kumar C.
        • Gnad F.
        • Nielsen M.L.
        • Rehman M.
        • Walther T.C.
        • Olsen J.V.
        • Mann M.
        Science. 2009; 325: 834-840
        • Naryzhny S.N.
        • Lee H.
        J. Biol. Chem. 2004; 279: 20194-20199
        • Muftuoglu M.
        • Kusumoto R.
        • Speina E.
        • Beck G.
        • Cheng W.H.
        • Bohr V.A.
        PLoS ONE. 2008; 3: e1918
        • Bhakat K.K.
        • Mokkapati S.K.
        • Boldogh I.
        • Hazra T.K.
        • Mitra S.
        Mol. Cell. Biol. 2006; 26: 1654-1665
        • Bhakat K.K.
        • Izumi T.
        • Yang S.H.
        • Hazra T.K.
        • Mitra S.
        EMBO J. 2003; 22: 6299-6309
        • Hasan S.
        • El-Andaloussi N.
        • Hardeland U.
        • Hassa P.O.
        • Bürki C.
        • Imhof R.
        • Schär P.
        • Hottiger M.O.
        Mol. Cell. 2002; 10: 1213-1222
        • Henneke G.
        • Koundrioukoff S.
        • Hübscher U.
        Oncogene. 2003; 22: 4301-4313
        • Guo Z.
        • Zheng L.
        • Xu H.
        • Dai H.
        • Zhou M.
        • Pascua M.R.
        • Chen Q.M.
        • Shen B.
        Nat. Chem. Biol. 2010; 6: 766-773
        • Budd M.E.
        • Campbell J.L.
        Proc. Natl. Acad. Sci. U.S.A. 1995; 92: 7642-7646
        • Pike J.E.
        • Henry R.A.
        • Burgers P.M.
        • Campbell J.L.
        • Bambara R.A.
        J. Biol. Chem. 2010; 285: 41712-41723
        • Ryu G.H.
        • Tanaka H.
        • Kim D.H.
        • Kim J.H.
        • Bae S.H.
        • Kwon Y.N.
        • Rhee J.S.
        • MacNeill S.A.
        • Seo Y.S.
        Nucleic Acids Res. 2004; 32: 4205-4216