Contacts between the 5′ Nuclease of DNA Polymerase I and Its DNA Substrate
2001; Elsevier BV; Volume: 276; Issue: 32 Linguagem: Inglês
10.1074/jbc.m100985200
ISSN1083-351X
AutoresYang Xu, Olga Potapova, Andres E. Leschziner, Nigel D. F. Grindley, Catherine M. Joyce,
Tópico(s)HIV/AIDS drug development and treatment
ResumoThe 5′ nuclease of DNA polymerase I (Pol I) of Escherichia coli is a member of an important class of prokaryotic and eukaryotic nucleases, involved in DNA replication and repair, with specificity for the junction between single-stranded and duplex DNA. We have investigated the interaction of the 5′ nuclease domain with DNA substrates from the standpoint of both the protein and the DNA. Phosphate ethylation interference showed that the nuclease binds to the nucleotides immediately surrounding the cleavage site and also contacts the complementary strand one-half turn away, indicating that contacts are made to one face only of the duplex portion of the DNA substrate. Phosphodiester contacts were investigated further using DNA substrates carrying unique methylphosphonate substitutions, together with mutations in the 5′ nuclease. These experiments suggested that two highly conserved basic residues, Lys78 and Arg81, are close to the phosphodiester immediately 5′ to the cleavage site, while a third highly conserved residue, Arg20, may interact with the phosphodiester 3′ to the cleavage site. Our results provide strong support for a DNA binding model proposed for the related exonuclease from bacteriophage T5, in which the conserved basic residues mentioned above define the two ends of a helical arch that forms part of the single-stranded DNA-binding region. The nine highly conserved carboxylates in the active site region appear to play a relatively minor role in substrate binding, although they are crucial for catalysis. In addition to binding the DNA backbone around the cleavage point, the 5′ nuclease also has a binding site for one or two frayed bases at the 3′ end of an upstream primer strand. In agreement with work in related systems, 5′ nuclease cleavage is blocked by duplex DNA in the 5′ tail, but the enzyme is quite tolerant of abasic DNA or polarity reversal within the 5′ tail. The 5′ nuclease of DNA polymerase I (Pol I) of Escherichia coli is a member of an important class of prokaryotic and eukaryotic nucleases, involved in DNA replication and repair, with specificity for the junction between single-stranded and duplex DNA. We have investigated the interaction of the 5′ nuclease domain with DNA substrates from the standpoint of both the protein and the DNA. Phosphate ethylation interference showed that the nuclease binds to the nucleotides immediately surrounding the cleavage site and also contacts the complementary strand one-half turn away, indicating that contacts are made to one face only of the duplex portion of the DNA substrate. Phosphodiester contacts were investigated further using DNA substrates carrying unique methylphosphonate substitutions, together with mutations in the 5′ nuclease. These experiments suggested that two highly conserved basic residues, Lys78 and Arg81, are close to the phosphodiester immediately 5′ to the cleavage site, while a third highly conserved residue, Arg20, may interact with the phosphodiester 3′ to the cleavage site. Our results provide strong support for a DNA binding model proposed for the related exonuclease from bacteriophage T5, in which the conserved basic residues mentioned above define the two ends of a helical arch that forms part of the single-stranded DNA-binding region. The nine highly conserved carboxylates in the active site region appear to play a relatively minor role in substrate binding, although they are crucial for catalysis. In addition to binding the DNA backbone around the cleavage point, the 5′ nuclease also has a binding site for one or two frayed bases at the 3′ end of an upstream primer strand. In agreement with work in related systems, 5′ nuclease cleavage is blocked by duplex DNA in the 5′ tail, but the enzyme is quite tolerant of abasic DNA or polarity reversal within the 5′ tail. DNA polymerase I flap endonuclease I DNA polymerase I (Pol I)1 of Escherichia coli has an intrinsic 5′ nuclease activity that is important for the removal of RNA primers from Okazaki fragments during lagging strand replication and for the removal of damaged nucleotides in DNA excision repair (1Kornberg A. Baker T.A. DNA Replication. 2nd Ed. W. H. Freeman and Co., San Francisco1992Google Scholar). Homologous 5′ nuclease domains are found in most other bacterial Pol I enzymes and in some bacteriophages, where the polymerase and 5′ nuclease exist as separate polypeptides (2Gutman P.D. Minton K.W. Nucleic Acids Res. 1993; 21: 4406-4407Crossref PubMed Scopus (54) Google Scholar, 3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar). The bacterial 5′ nuclease family also shows substantial sequence similarity to the FEN-1 eukaryotic nucleases, which are involved in various DNA transactions (4Robins P. Pappin D.J.C. Wood R.D. Lindahl T. J. Biol. Chem. 1994; 269: 28535-28538Abstract Full Text PDF PubMed Google Scholar, 5Lieber M.R. BioEssays. 1997; 19: 233-240Crossref PubMed Scopus (396) Google Scholar, 6Mueser T.C. Nossal N.G. Hyde C.C. Cell. 1996; 85: 1101-1112Abstract Full Text Full Text PDF PubMed Scopus (163) Google Scholar, 7Harrington J.J. Lieber M.R. Genes Dev. 1994; 8: 1344-1355Crossref PubMed Scopus (256) Google Scholar, 8Kim K. Biade S. Matsumoto Y. J. Biol. Chem. 1998; 273: 8842-8848Abstract Full Text Full Text PDF PubMed Scopus (188) Google Scholar). Although the 5′ nucleases were originally called 5′-3′ exonucleases, these enzymes are more accurately described as structure-specific nucleases, with specificity for the junction between a base paired region and a single-stranded 5′ overhang or "flap" (9Lundquist R. Olivera B. Cell. 1982; 31: 53-60Abstract Full Text PDF PubMed Scopus (57) Google Scholar, 10Lyamichev V. Brow M.A.D. Dahlberg J.E. Science. 1993; 260: 778-783Crossref PubMed Scopus (306) Google Scholar, 11Harrington J.J. Lieber M.R. EMBO J. 1994; 13: 1235-1246Crossref PubMed Scopus (373) Google Scholar). Cleavage by these nucleases usually takes place between the first two paired bases at the junction between the duplex and the single-stranded 5′ tail (see Fig. 1), although some variability has been observed (10Lyamichev V. Brow M.A.D. Dahlberg J.E. Science. 1993; 260: 778-783Crossref PubMed Scopus (306) Google Scholar, 11Harrington J.J. Lieber M.R. EMBO J. 1994; 13: 1235-1246Crossref PubMed Scopus (373) Google Scholar, 12Bhagwat M. Hobbs L.J. Nossal N.G. J. Biol. Chem. 1997; 272: 28523-28530Abstract Full Text Full Text PDF PubMed Scopus (21) Google Scholar, 13Murante R.S. Rumbaugh J.A. Barnes C.J. Norton J.R. Bambara R.A. J. Biol. Chem. 1996; 271: 25888-25897Abstract Full Text Full Text PDF PubMed Scopus (55) Google Scholar). Recent work suggests that the preferred substrate of the bacterial nucleases has a single unpaired base at the 3′ end of the primer upstream of the 5′ nuclease cleavage site (thus facilitating formation of a ligatable nick), and it seems likely that any variability in the observed cleavage position may reflect rearrangement via branch migration of the 5′ and 3′ single-stranded flaps (14Lyamichev V. Brow M.A.D. Varvel V.E. Dahlberg J.E. Proc. Natl. Acad. Sci. U. S. A. 1999; 96: 6143-6148Crossref PubMed Scopus (64) Google Scholar, 15Xu Y. Grindley N.D.F. Joyce C.M. J. Biol. Chem. 2000; 275: 20949-20955Abstract Full Text Full Text PDF PubMed Scopus (37) Google Scholar). Alignment of the sequences of the bacterial and bacteriophage 5′ nucleases revealed nine invariant carboxylates (2Gutman P.D. Minton K.W. Nucleic Acids Res. 1993; 21: 4406-4407Crossref PubMed Scopus (54) Google Scholar, 3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar), the majority of which are also conserved in the FEN-1 family (6Mueser T.C. Nossal N.G. Hyde C.C. Cell. 1996; 85: 1101-1112Abstract Full Text Full Text PDF PubMed Scopus (163) Google Scholar, 16Shen B. Qiu J. Hosfield D. Tainer J.A. Trends Biochem. Sci. 1998; 23: 171-173Abstract Full Text Full Text PDF PubMed Scopus (62) Google Scholar). The large number of conserved carboxylates raises the possibility that the 5′ nuclease reaction, like the polymerase and 3′-5′ exonuclease reactions, may be catalyzed by divalent metal ions coordinated to the enzyme via carboxylate ligands (17Joyce C.M. Steitz T.A. Annu. Rev. Biochem. 1994; 63: 777-822Crossref PubMed Scopus (570) Google Scholar). Moreover, mutagenesis studies in several systems have demonstrated the importance of the carboxylates in the 5′ nuclease reaction (3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar, 18Mizrahi V. Huberts P. Nucleic Acids Res. 1996; 24: 4845-4852Crossref PubMed Scopus (37) Google Scholar, 19Bhagwat M. Meara D. Nossal N.G. J. Biol. Chem. 1997; 272: 28531-28538Abstract Full Text Full Text PDF PubMed Scopus (34) Google Scholar, 20Shen B. Nolan J.P. Sklar L.A. Park M.S. J. Biol. Chem. 1996; 271: 9173-9176Abstract Full Text Full Text PDF PubMed Scopus (101) Google Scholar, 21Shen B. Nolan J.P. Sklar L.A. Park M.S. Nucleic Acids Res. 1997; 25: 3332-3338Crossref PubMed Scopus (95) Google Scholar). Crystal structures have been determined for three of the prokaryotic 5′ nucleases, the intrinsic 5′ nuclease of Thermus aquaticus(Taq) DNA polymerase I (22Kim Y. Eom S.H. Wang J. Lee D.S. Suh S.W. Steitz T.A. Nature. 1995; 376: 612-616Crossref PubMed Scopus (329) Google Scholar), bacteriophage T4 RNase H (6Mueser T.C. Nossal N.G. Hyde C.C. Cell. 1996; 85: 1101-1112Abstract Full Text Full Text PDF PubMed Scopus (163) Google Scholar), and bacteriophage T5 exonuclease (23Ceska T.A. Sayers J.R. Stier G. Sück D. Nature. 1996; 382: 90-93Crossref PubMed Scopus (165) Google Scholar), and for two archaebacterial FEN-1 analogs (24Hwang K.Y. Baek K. Kim H.-Y. Cho Y. Nat. Struct. Biol. 1998; 5: 707-713Crossref PubMed Scopus (149) Google Scholar, 25Hosfield D.J. Mol C.D. Shen B. Tainer J.A. Cell. 1998; 95: 135-146Abstract Full Text Full Text PDF PubMed Scopus (225) Google Scholar). The core structures of all five enzymes are very similar with the invariant carboxylates clustered in the central portion of the molecule, some coordinated to divalent metal ions, although the precise locations of the metal ions and the ligand geometries are somewhat variable (3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar, 26Sayers J.R. Artymiuk P.J. Nat. Struct. Biol. 1998; 5: 668-670Crossref PubMed Scopus (17) Google Scholar). In the absence of any crystalline complexes with bound DNA, little information exists on how the 5′ nucleases interact with their substrate. A plausible candidate for a DNA-binding region in the prokaryotic 5′ nucleases is a cluster of highly conserved and mostly basic residues. The relevant region of the protein structure is not well resolved in the Taq DNA polymerase and T4 RNase H structures, but in T5 exonuclease it is located on one side of an unusual helical arch structure (23Ceska T.A. Sayers J.R. Stier G. Sück D. Nature. 1996; 382: 90-93Crossref PubMed Scopus (165) Google Scholar). The two FEN-1 structures have a flexible loop in place of the helical arch (24Hwang K.Y. Baek K. Kim H.-Y. Cho Y. Nat. Struct. Biol. 1998; 5: 707-713Crossref PubMed Scopus (149) Google Scholar, 25Hosfield D.J. Mol C.D. Shen B. Tainer J.A. Cell. 1998; 95: 135-146Abstract Full Text Full Text PDF PubMed Scopus (225) Google Scholar). In all three structures the loop or arch is large enough to accommodate single-stranded DNA, suggesting that the single-stranded 5′ end of the DNA substrate may be threaded through this part of the protein (Refs.23Ceska T.A. Sayers J.R. Stier G. Sück D. Nature. 1996; 382: 90-93Crossref PubMed Scopus (165) Google Scholar, 24Hwang K.Y. Baek K. Kim H.-Y. Cho Y. Nat. Struct. Biol. 1998; 5: 707-713Crossref PubMed Scopus (149) Google Scholar, and 26Sayers J.R. Artymiuk P.J. Nat. Struct. Biol. 1998; 5: 668-670Crossref PubMed Scopus (17) Google Scholar, see Fig. 2 B), with the duplex DNA portion held close to the base of the loop or arch by a helix-loop-helix motif present in all five of the available structures (25Hosfield D.J. Mol C.D. Shen B. Tainer J.A. Cell. 1998; 95: 135-146Abstract Full Text Full Text PDF PubMed Scopus (225) Google Scholar). The threading model was originally proposed based on observations that 5′ nuclease cleavage requires a free 5′ end and is blocked by significant obstructions (e.g. annealed primers) in the single-stranded tail (3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar, 10Lyamichev V. Brow M.A.D. Dahlberg J.E. Science. 1993; 260: 778-783Crossref PubMed Scopus (306) Google Scholar, 27Murante R.S. Rust L. Bambara R.A. J. Biol. Chem. 1995; 270: 30377-30383Abstract Full Text Full Text PDF PubMed Scopus (186) Google Scholar). However, recent work has shown that FEN-1 can tolerate a variety of modifications, including an 11-nucleotide branch, within the 5′ flap DNA, prompting re-examination of the threading model (28Bornarth C.J. Ranalli T.A. Henricksen L.A. Wahl A.F. Bambara R.A. Biochemistry. 1999; 38: 13347-13354Crossref PubMed Scopus (69) Google Scholar). In this work, we have investigated the interaction between the 5′ nuclease and its DNA substrate from the perspective of both the protein and the DNA, and have attempted to identify contacts between particular protein side chains and individual phosphodiester groups on the DNA. Oligonucleotides for site-directed mutagenesis and for 5′ nuclease substrates were synthesized by the Keck Biotechnology Resource Laboratory at Yale Medical School. Those used as reaction substrates were purified by gel electrophoresis. Oligonucleotide concentrations were determined spectrophotometrically using calculated extinction coefficients (29Puglisi J.D. Tinoco I. Methods Enzymol. 1989; 180: 304-325Crossref PubMed Scopus (662) Google Scholar). DNA oligonucleotides containing abasic spacers or regions with reversed polarity were kindly provided by Dr. Daniel Kaplan, and have been described elsewhere (30Kaplan D.L. J. Mol. Biol. 2000; 301: 285-299Crossref PubMed Scopus (108) Google Scholar). Radiolabeled nucleotides were from Amersham Pharmacia Biotech. N-Ethyl-N-nitrosourea was purchased from Sigma. DNase I was from Cooper Biomedical. T4 polynucleotide kinase, DNA ligase, and restriction endonucleases were from New England Biolabs or Roche Molecular Biochemicals and were used according to the accompanying instructions. Following our published methods (3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar), mutations in the 5′ nuclease were constructed and subcloned into an expression plasmid for the 323-amino acid 5′ nuclease domain, and the mutant proteins were purified by fast protein liquid chromatography (Amersham Pharmacia Biotech). To remove low levels of contamination by intact Pol I, the purification method was modified in either of two ways. When preparing 5′ nuclease mutants on a small scale, the peak fractions from the MonoQ HR 16/20 column were assayed for polymerase activity (31Setlow P. Methods Enzymol. 1974; 29: 3-12Crossref PubMed Scopus (36) Google Scholar), and only those fractions that had no measurable polymerase activity were combined and applied to the phenyl-Superose column. For large scale purification, a final gel filtration step was included. The 5′ nuclease pool from the phenyl-Superose column was concentrated by ammonium sulfate precipitation, applied to a HiLoad 16/60 Superdex 200 prep grade column (120-ml bed volume) and eluted with 50 mm Tris-HCl, pH 7.5, 1 mm dithiothreitol, 100 mm NaCl. These modifications were necessary for the present study because low-level contamination by full-length Pol I interfered with DNase I footprinting experiments, particularly with weak binding 5′ nuclease mutants. Because the DNA binding affinity of the polymerase domain is much greater than that of the 5′ nuclease, competition between the two was a problem even though the amount of Pol I was not sufficient to affect enzyme activity assays and was not detectable on a Coomassie Blue-stained gel. We believe that the contaminating Pol I in our preparations was derived from the chromosomal polA locus rather than from readthrough of the amber codon in our expression plasmid for the 5′ nuclease domain (3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar) because the problem was not alleviated by subcloning the 5′ nuclease mutations into an analogous expression construct that lacked the complete polymerase coding region. Single-turnover measurements of 5′ nuclease cleavage were carried out using the substrates shown in Fig.1 a. The DNA oligonucleotides were labeled with32P at either the 5′ or the 3′ end (see legend to Fig. 1). Reactions contained ≈5 nm DNA substrate and the 5′ nuclease at a series of 7–8 concentrations (typically from 0.1 to 40 µm), chosen so as to bracket the K Dvalue. All reactions were carried out at ambient temperature (23 °C) in a buffer containing 50 mm Tris-HCl, pH 7.6, 5 mm MgCl2, 100 mm NaCl, and 5 µm p(dT)10. The reactions catalyzed by the mutant 5′ nuclease derivatives (requiring sampling at time intervals of ≥10 s) were conducted as described previously (3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar). The reaction catalyzed by the wild-type 5′ nuclease was too fast for manual sampling, and was instead carried out on a rapid quench-flow instrument (KinTek Corp., Model RQF-3) by mixing a solution containing the DNA substrate with an equal volume of enzyme solution (both at twice the desired final concentration in the buffer described above) and quenching with 150 mm EDTA. Samples, obtained either manually or from rapid-quench experiments, were fractionated over a denaturing 10% polyacrylamide gel, and quantitated as described previously (3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar). For each set of reaction conditions, the enzyme concentration was much higher than that of the substrate, so that the reaction followed pseudo first-order kinetics. The observed rate constants (k obs) were then plotted as a function of enzyme concentration (E 0), and the DNA binding constant, K D, and cleavage rate constant,k c, were obtained by fitting to the equation:k obs =k c E 0/(K D +E 0). All determinations were carried out at least in duplicate. Details of the reaction conditions for 5′ nuclease cleavage of modified substrates are given in the relevant figure legends. Footprinting of the 112-mer double-hairpin oligonucleotide (Fig. 1 b), as a function of 5′ nuclease concentration, was carried out as described previously (15Xu Y. Grindley N.D.F. Joyce C.M. J. Biol. Chem. 2000; 275: 20949-20955Abstract Full Text Full Text PDF PubMed Scopus (37) Google Scholar, 32Polesky A.H. Steitz T.A. Grindley N.D.F. Joyce C.M. J. Biol. Chem. 1990; 265: 14579-14591Abstract Full Text PDF PubMed Google Scholar). After gel fractionation, appropriate fragments were quantitated by phosphorimaging. For each enzyme concentration, the percent of uncomplexed DNA was calculated by determining the ratio of the radioactivity in a DNA band within the protected region to that in a band outside the protected region, and comparing with the same ratio from a control lane with no enzyme present. The binding constant,K D, equals the enzyme concentration at which 50% of the DNA is bound, which was determined by curve fitting. For this analysis to be valid, the DNA concentration in the reaction must be substantially below K D, so that the total enzyme concentration approximates the concentration of free enzyme. This condition was fulfilled because of the relatively highK D values for the 5′ nuclease and its mutant derivatives. CD spectra were recorded at 20 °C on a solution of wild-type or mutant 5′ nuclease (≈4–10 µmin 20 mm potassium phosphate, pH 7.0, 15 mmNaCl, 15% (v/v) glycerol) in a 2-mm path length quartz cuvette (110-QS, Hellma) which was placed in a thermostated sample holder in an Aviv 62 DS CD spectrometer (Lakewood, NJ). Thermal denaturation measurements were performed in the same buffer by monitoring the ellipticity at 222 nm as a function of temperature, over the range 10–70 °C in 1 °C increments (samples were equilibrated for 2 min at each temperature). The data were processed as described (33Astatke M. Grindley N.D.F. Joyce C.M. J. Biol. Chem. 1995; 270: 1945-1954Abstract Full Text Full Text PDF PubMed Scopus (111) Google Scholar), to give T m, the temperature at which the protein is 50% unfolded. The procedure was modified from published procedures (34Rimphanitchayakit V. Grindley N.D.F. Jost J.P. Saluz H.P. A Laboratory Guide to in Vitro Studies of Protein-DNA Interactions. Birkhauser Verlag Basel, Basel1991: 111-120Crossref Google Scholar, 35Wissman A. Hillen W. Methods Enzymol. 1991; 208: 365-379Crossref PubMed Scopus (62) Google Scholar). The 5′ end-labeled 132-mer double-hairpin substrate (Fig. 1 b) was used to analyze phosphate contacts close to the cleavage site. To ≈2 pmol of the DNA substrate in 0.1 ml of 50 mm sodium cacodylate, pH 8, was added 0.1 ml of ethanol saturated with ethylnitrosourea at 50 °C. After incubation at 50 °C for about 25 min, the ethylnitrosourea was removed by five extractions with 1-ml portions of water-saturated ether. The ethylated DNA solution (≈0.1 ml) was adjusted to 50 mm Tris, pH 7.5, 5 mm MgCl2, and incubated at room temperature with wild-type 5′ nuclease (≈200 pmol). One-third of the reaction mixture was quenched with an equal volume of formamide- dyes containing 30 mm EDTA, at time intervals chosen so as to obtain three samples having ≈25, 50, and 75% product formation, respectively. The 36-mer product was separated from unreacted 132-mer substrate in each sample by electrophoresis on an 8% polyacrylamide gel containing 40% (v/v) formamide and 7 murea. The DNA bands were located by autoradiography, excised, eluted into 240 µl of 10 mm Tris-HCl, pH 8.0, 0.1 mmEDTA, containing 0.5 m NH4OAc, and concentrated by ethanol precipitation. The resulting DNA samples were cleaved at the ethylated phosphate positions by heating for 30 min at 90 °C in 10 µl of 10 mm sodium phosphate, pH 8.0, 1 mmEDTA, containing 75 mm NaOH. A portion (up to 4 µl) of each sample was mixed with 2 µl of formamide- dyes, and fractionated on a 12% polyacrylamide-urea gel. Size markers were generated by restriction enzyme digestion (see Fig. 1 b), and by limited chemical cleavage at guanine residues (36Maxam A.M. Gilbert W. Proc. Natl. Acad. Sci. U. S. A. 1977; 74: 560-564Crossref PubMed Scopus (5477) Google Scholar). Data analysis is described in the legend to Fig. 3. To analyze phosphate contacts on the DNA strand opposite the cleavage site, the same procedure was carried out using the 112-mer, and analyzing samples from the pool of unreacted substrate on an 8% polyacrylamide gel containing 40% (v/v) formamide and 7 m urea. To determine which amino acid side chains of the 5′ nuclease are involved in binding the DNA substrate, we studied mutations in two groups of highly conserved residues (Table I). One was the nine carboxylates, shown to be crucial for 5′ nuclease cleavage in our previous study (3Xu Y. Derbyshire V. Ng K. Sun X.C. Grindley N.D.F. Joyce C.M. J. Mol. Biol. 1997; 268: 284-302Crossref PubMed Scopus (51) Google Scholar). We also made alanine substitutions at Arg20, Arg70, Tyr77, Lys78, and Arg81 (Fig. 2 A); in the structure of the T5 5′ nuclease the equivalent residues define the base of the helical arch that is proposed to form part of the DNA-binding site (23Ceska T.A. Sayers J.R. Stier G. Sück D. Nature. 1996; 382: 90-93Crossref PubMed Scopus (165) Google Scholar) (Fig. 2 B). The effects of all these mutations were studied in context of the separate 5′ nuclease domain (residues 1–323 of Pol I) so as to avoid ambiguities due to the DNA binding properties of the other domains of Pol I.Table IConservation of 5′ nuclease residues mutated in this workResidue inE. coli Pol IConservation1-aThe alignment from which this analysis was derived is available on the web at pantheon.yale.edu/∼cjoyce/align.html.47 Bacterial pols11 Bacterial nucleases1-bCoding regions identified as members of the 5′ nuclease family on the basis of sequence similarity. In a few cases, e.g. Ref. 37, nuclease activity has been demonstrated.4 Phage nucleasesAsp1347 Asp11 Asp4 AspArg2047 Arg10 Arg, 1 Ala1 Arg, 2 Gln, 1 Ser1-cDeduced by structural alignment of T4 and T5 5′ nucleases.Asp6346 Asp, 1 Glu11 Asp4 AspArg7047 Arg11 Arg4 ArgTyr7747 Tyr11 Tyr4 TyrLys7847 Lys11 Lys4 LysArg8147 Arg11 Arg4 ArgGlu11347 Glu11 Glu4 GluAsp11547 Asp11 Asp4 AspAsp11647 Asp11 Asp4 AspAsp13847 Asp11 Asp4 AspAsp14047 Asp9 Asp, 2 Gly4 AspAsp18547 Asp9 Asp, 1 Ile, 1 Val4 AspAsp18847 Asp9 Asp, 2 Ser4 Asp1-a The alignment from which this analysis was derived is available on the web at pantheon.yale.edu/∼cjoyce/align.html.1-b Coding regions identified as members of the 5′ nuclease family on the basis of sequence similarity. In a few cases, e.g. Ref. 37Shafritz K.M. Sandigursky M. Franklin W.A. Nucleic Acids Res. 1998; 26: 2593-2597Crossref PubMed Scopus (16) Google Scholar, nuclease activity has been demonstrated.1-c Deduced by structural alignment of T4 and T5 5′ nucleases. Open table in a new tab Cleavage of our standard (22 + 68)-mer substrate (Fig. 1) by the wild-type and mutant 5′ nuclease derivatives was measured under single-turnover conditions (excess enzyme) as a function of enzyme concentration, giving the enzyme-DNA dissociation constant (K D) and the maximum cleavage rate (k c), which reflects the rate of steps up to and including phosphoryl transfer (Table II). All of the mutations tested caused substantial decreases in cleavage rate, ranging from ≈20-fold (R20A and R81A) to >5 × 105-fold (D115A). R20A and K78A were the only mutations that caused a significant decrease in DNA binding affinity; some of the carboxylate mutations, particularly D63A, D138A, and D185A, caused an increase in DNA binding affinity. For a few 5′ nuclease mutants, the DNA binding affinity was measured by a DNase I footprint titration; this showed good agreement with the kinetic determinations and had the added advantage that the measured binding constant is related to a binding site which can be visualized on a gel (Ref. 15Xu Y. Grindley N.D.F. Joyce C.M. J. Biol. Chem. 2000; 275: 20949-20955Abstract Full Text Full Text PDF PubMed Scopus (37) Google Scholar gives details of the 5′ nuclease footprint). Footprinting was clearly the preferred method for the D115A mutant protein, which had no measurable activity in the 5′ nuclease assay; conversely, footprinting was not appropriate for mutants with relatively high nuclease activity which degraded the DNA substrate during the experiment.Table IIProperties of wild-type and mutant derivatives of the 5′ nuclease domainEnzymeKinetics2-aKinetic measurements were made using the (22 + 68)-mer substrate (Fig. 1 a).Footprinting2-bDNase I protection was carried out using the 112-mer substrate (Fig. 1 b). Small differences in binding affinity between the two substrates used inK D measurements are to be expected because of their different sequences, and because the (22 + 68)-mer has a nick and the 112-mer has a single base gap upstream of the cleavage site (see also Ref. 15).T m(°C)k cK DK DNo Mg2+5 mmMg2+2-cFor the three proteins tested, the CD spectra in the presence and absence of Mg2+ were superimposable.min−1µmµmWT47 ± 12-dUse of the rapid-quench-flow instrument gave a more accurate measurement of the reaction rate for the wild-type 5′-3′ exonuclease, and indicated that this rate is about 10-fold faster than that reported in our previous study.4.7 ± 0.7 (2)2-cFor the three proteins tested, the CD spectra in the presence and absence of Mg2+ were superimposable.41 ± 1 (2)49R20A2.8 ± 0.827 ± 15 (3)ND2-fND, not determined; however, the high level of 5′ nuclease activity argues against any significant alteration to the structure.R70A(4.3 ± 0.4) × 10−24.0 ± 0.9 (2)39Y77A(4.0 ± 0.5) × 10−34.8 ± 0.6 (2)1.8 ± 0.1 (2)42K78A(4.3 ± 1.1) × 10−211 ± 2 (2)26 ± 3 (2)44R81A2.2 ± 0.15.5 ± 1.5 (4)40D13N(7.5 ± 0.9) × 10−47.6 ± 1.4 (3)44D63A(2.0 ± 0.4) × 10−20.26 ± 0.16 (3)38E113A(1.4 ± 0.4) × 10−22.7 ± 2.0 (5)45D115A<1 × 10−42-gThe reaction was too slow to measure.1.2 ± 0.1 (2)46 ± 1 (2)54D116A(6.4 ± 1.6) × 10−43.0 ± 1.2 (3)43D138N(9.4 ± 1.9) × 10−40.30 ± 0.17 (3)39D140A(5.0 ± 1.0) × 10−44.6 ± 2.1 (3)50 ± 1 (2)55D185A(9.7 ± 1.6) × 10−30.18 ± 0.13 (3)0.16 ± 0.02 (2)37D188A(5.3 ± 1.0) × 10−31.4 ± 1.0 (4)412-a Kinetic measurements were made using the (22 + 68)-mer substrate (Fig. 1 a).2-b DNase I protection was carried out using the 112-mer substrate (Fig. 1 b). Small differences in binding affinity between the two substrates used inK D measurements are to be expected because of their different sequences, and because the (22 + 68)-mer has a nick and the 112-mer has a single base gap upstream of the cleavage site (see also Ref. 15Xu Y. Grindley N.D.F. Joyce C.M. J. Biol. Chem. 2000; 275: 20949-20955Abstract Full Text Full Text PDF PubMed Scopus (37) Google Scholar).2-c For the three proteins tested, the CD spectra in the presence and absence of Mg2+ were superimposable.2-d Use of the rapid-quench-flow instrument gave a more accurate measurement of the reaction rate for the wild-type 5′-3′ exonuclease, and indicated that this rate is about 10-fold faster than that reported in our
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