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Structural Requirements at the Catalytic Site of the Heteroduplex Substrate for Human RNase H1 Catalysis

2004; Elsevier BV; Volume: 279; Issue: 35 Linguagem: Inglês

10.1074/jbc.m405035200

1083-351X

Walt F. Lima, Josh G. Nichols, Hongjiang Wu, Thazha P. Prakash, M.T. Migawa, Tadensz K. Wyrzykiewicz, Balkrishen Bhat, Stanley T. Crooke,

Bacterial Genetics and Biotechnology

Human RNase H1 cleaves RNA exclusively in an RNA/DNA duplex; neither double-strand DNA nor double-strand RNA is a viable substrate. Previous studies suggest that the helical geometry and sugar conformation of the DNA and RNA may play a role in the selective recognition of the heteroduplex substrate by the enzyme. We systematically evaluated the influence of sugar conformation, minor groove bulk, and conformational flexibility of the heteroduplex on enzyme efficiency. Modified nucleotides were introduced into the oligodeoxyribonucleotide at the catalytic site of the heteroduplex and consisted of southern, northern, and eastern biased sugars with and without 2′-substituents, non-hydrogen bonding base modifications, abasic deoxyribonucleotides, intranucleotide hydrocarbon linkers, and a ganciclovir-modified deoxyribonucleotide. Heteroduplexes containing modifications exhibiting strong northern or southern conformational biases with and without a bulky 2′-substituent were cleaved at a significantly slower rate than the unmodified substrate. Modifications imparting the greatest degree of conformational flexibility were the poorest substrates, resulting in dramatically slower cleavage rates for the ribonucleotide opposing the modification and the surrounding ribonucleotides. Finally, modified heteroduplexes containing modifications predicted to mimic the sugar pucker and conformational flexibility of the deoxyribonucleotide exhibited cleavage rates comparable with those of the unmodified substrate. These data suggest that sugar conformation, minor groove width, and the relative positions of the intra- and internucleotide phosphates are the crucial determinants in the selective recognition of the heteroduplex substrate by human RNase H1 and offer immediate steps to improve the performance of DNA-like antisense oligonucleotides. Human RNase H1 cleaves RNA exclusively in an RNA/DNA duplex; neither double-strand DNA nor double-strand RNA is a viable substrate. Previous studies suggest that the helical geometry and sugar conformation of the DNA and RNA may play a role in the selective recognition of the heteroduplex substrate by the enzyme. We systematically evaluated the influence of sugar conformation, minor groove bulk, and conformational flexibility of the heteroduplex on enzyme efficiency. Modified nucleotides were introduced into the oligodeoxyribonucleotide at the catalytic site of the heteroduplex and consisted of southern, northern, and eastern biased sugars with and without 2′-substituents, non-hydrogen bonding base modifications, abasic deoxyribonucleotides, intranucleotide hydrocarbon linkers, and a ganciclovir-modified deoxyribonucleotide. Heteroduplexes containing modifications exhibiting strong northern or southern conformational biases with and without a bulky 2′-substituent were cleaved at a significantly slower rate than the unmodified substrate. Modifications imparting the greatest degree of conformational flexibility were the poorest substrates, resulting in dramatically slower cleavage rates for the ribonucleotide opposing the modification and the surrounding ribonucleotides. Finally, modified heteroduplexes containing modifications predicted to mimic the sugar pucker and conformational flexibility of the deoxyribonucleotide exhibited cleavage rates comparable with those of the unmodified substrate. These data suggest that sugar conformation, minor groove width, and the relative positions of the intra- and internucleotide phosphates are the crucial determinants in the selective recognition of the heteroduplex substrate by human RNase H1 and offer immediate steps to improve the performance of DNA-like antisense oligonucleotides. RNase H hydrolyzes RNA in RNA-DNA hybrids (1Stein H. Hausen P. Science. 1969; 166: 393-395Crossref PubMed Scopus (183) Google Scholar). RNase H activity appears to be ubiquitous in eukaryotes and bacteria (2Itaya M. Kondo K. Nucleic Acids Res. 1991; 19: 4443-4449Crossref PubMed Scopus (63) Google Scholar, 3Itaya M. McKelvin D. Chatterjie S.K. Crouch R.J. Mol. Gen. Genet. 1991; 227: 438-445Crossref PubMed Scopus (55) Google Scholar, 4Kanaya S. Itaya M. J. Biol. Chem. 1992; 267: 10184-10192Abstract Full Text PDF PubMed Google Scholar, 5Busen W. J. Biol. Chem. 1980; 255: 9434-9443Abstract Full Text PDF PubMed Google Scholar, 6Rong Y.W. Carl P.L. Biochemistry. 1990; 29: 383-389Crossref PubMed Scopus (34) Google Scholar, 7Eder P.S. Walder R.T. Walder J.A. Biochimie (Paris). 1993; 75: 123-126Crossref PubMed Scopus (104) Google Scholar). Although RNases H constitute a family of proteins of varying molecular mass, the nucleolytic activity and substrate requirements appear to be similar for the various isotypes. For example, all RNases H studied to date function as endonucleases, exhibiting limited sequence specificity and requiring divalent cations (e.g. Mg2+ and Mn2+) to produce cleavage products with 5′-phosphate and 3′-hydroxyl termini (8Crouch R.J. Dirksen M.L. Linn S.M. Roberts R.J. Nucleases. Cold Spring Harbor Laboratory Press, Plainview, NY1982: 211-241Google Scholar). Recently, two human RNase H genes have been cloned and expressed (9Wu H. Lima W.F. Crooke S.T. Antisense Nucleic Acid Drug Dev. 1998; 8: 53-61Crossref PubMed Scopus (55) Google Scholar, 10Busen W. Peters J.H. Hausen P. Eur. J. Biochem. 1977; 74: 203-208Crossref PubMed Scopus (49) Google Scholar, 11Turchi J.J. Huang L. Murante R.S. Kim Y. Bambara R.A. Proc. Natl. Acad. Sci. U. S. A. 1994; 91: 9803-9807Crossref PubMed Scopus (169) Google Scholar). RNase H1 is a 286-amino acid protein and is expressed ubiquitously in human cells and tissues (9Wu H. Lima W.F. Crooke S.T. Antisense Nucleic Acid Drug Dev. 1998; 8: 53-61Crossref PubMed Scopus (55) Google Scholar). The amino acid sequence of human RNase H1 displays strong homology to RNase H1 from yeast, chicken, Escherichia coli, and mouse (9Wu H. Lima W.F. Crooke S.T. Antisense Nucleic Acid Drug Dev. 1998; 8: 53-61Crossref PubMed Scopus (55) Google Scholar). The human RNase H2 enzyme is a 299-amino acid protein with a calculated mass of 33.4 kDa and has also been shown to be ubiquitously expressed in human cells and tissues (10Busen W. Peters J.H. Hausen P. Eur. J. Biochem. 1977; 74: 203-208Crossref PubMed Scopus (49) Google Scholar). 1H. Wu, unpublished data. Human RNase H2 shares strong amino acid sequence homology with RNase H2 from Caenorhabditis elegans, yeast, and E. coli (10Busen W. Peters J.H. Hausen P. Eur. J. Biochem. 1977; 74: 203-208Crossref PubMed Scopus (49) Google Scholar). Although the biological roles for the human enzymes are not fully understood, RNase H2 appears to be involved in de novo DNA replication, and RNase H1 has been shown in mice to be important for mitochondrial DNA replication (12Ceritelli S.M. Frolova E.G. Feng C. Grinberg A. Love P.E. Crouch R.J. Mol. Cell. 2003; 11: 807-815Abstract Full Text Full Text PDF PubMed Scopus (264) Google Scholar). Human RNase H1 has been shown to play a dominant role in the activity of DNA-like antisense oligonucleotides (13Wu H. Lima W.F. Zhang H. Fan A. Sun H. Crooke S.T. J. Biol. Chem. 2004; 279: 17181-17189Abstract Full Text Full Text PDF PubMed Scopus (241) Google Scholar). Human RNase H1 protein is overexpressed in both several cell lines and mouse liver, and the level of human RNase H1 is reduced by using DNA-like antisense oligonucleotides (ASOs) 2The abbreviations used are: ASOs, antisense oligonucleotides; HPLC, high performance liquid chromatography. and small interfering RNAs targeting the enzyme. The effects of these manipulations on the potencies of a number of DNA-like ASOs to several different target RNAs showed that increasing the level and activity of human RNase H1 increases the potency of the ASOs (13Wu H. Lima W.F. Zhang H. Fan A. Sun H. Crooke S.T. J. Biol. Chem. 2004; 279: 17181-17189Abstract Full Text Full Text PDF PubMed Scopus (241) Google Scholar). Moreover, overexpression of human RNase H1 in mouse liver increases the potency of a DNA-like ASO targeting Fas after intravenous administration. Finally, reducing the level and activity of RNase H1 reduces the potencies of the ASOs (13Wu H. Lima W.F. Zhang H. Fan A. Sun H. Crooke S.T. J. Biol. Chem. 2004; 279: 17181-17189Abstract Full Text Full Text PDF PubMed Scopus (241) Google Scholar). The structure of human RNase H1 was shown to consist of a 73-amino acid region homologous to the RNA-binding domain of yeast RNase H1 at the N terminus of the protein and separated from the catalytic domain by a 62-amino acid spacer region (14Wu H. Lima W.F. Crooke S.T. J. Biol. Chem. 2001; 276: 23547-23553Abstract Full Text Full Text PDF PubMed Scopus (66) Google Scholar, 15Cerritelli S.M. Crouch R.J. RNA (N. Y.). 1995; 1: 246-259PubMed Google Scholar, 16Evans S.P. Bycroft M. J. Mol. Biol. 1999; 291: 661-669Crossref PubMed Scopus (30) Google Scholar). The catalytic domain is highly conserved in the amino acid sequences of other RNase H1 proteins and contains the key catalytic and substrate-binding residues required for activity (14Wu H. Lima W.F. Crooke S.T. J. Biol. Chem. 2001; 276: 23547-23553Abstract Full Text Full Text PDF PubMed Scopus (66) Google Scholar, 17Kanaya S. Katsuda-Kakai C. Ikehara M. J. Biol. Chem. 1991; 266: 11621-11627Abstract Full Text PDF PubMed Google Scholar, 18Nakamura H. Oda Y. Iwai S. Inoue H. Ohtsuka E. Kanaya S. Kimura S. Katsuda C. Katayanagi K. Morikawa K. Miyashiro H. Ikehara M. Proc. Natl. Acad. Sci. U. S. A. 1991; 88: 11535-11539Crossref PubMed Scopus (196) Google Scholar, 19Katayanagi K. Miyagawa M. Matsushima M. Ishkiawa M. Kanaya S. Ikehara M. Matsuzaki T. Morikawa K. Nature. 1990; 347: 306-309Crossref PubMed Scopus (308) Google Scholar, 20Yang W. Hendrickson W.A. Crouch R.J. Satow Y. Science. 1990; 249: 1398-1405Crossref PubMed Scopus (453) Google Scholar). Site-directed mutagenesis of human RNase H1 revealed that the spacer region is required for RNase H activity (14Wu H. Lima W.F. Crooke S.T. J. Biol. Chem. 2001; 276: 23547-23553Abstract Full Text Full Text PDF PubMed Scopus (66) Google Scholar). Although the RNA-binding domain was shown not to be required for RNase H activity, this region is responsible for the enhanced binding affinity of the human enzyme for the heteroduplex substrate as well as the strong positional preference for cleavage exhibited by the enzyme (14Wu H. Lima W.F. Crooke S.T. J. Biol. Chem. 2001; 276: 23547-23553Abstract Full Text Full Text PDF PubMed Scopus (66) Google Scholar, 21Lima W.F. Wu H. Nichols J. Prakash T.P. Ravikumar V. Crooke S.T. J. Biol. Chem. 2003; 278: 49860-49867Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar). The RNA-binding domain of human RNase H1 is conserved in other eukaryotic RNases H1, and the highly conserved lysines at positions 59 and 60 of human RNase H1 have been shown to be important for binding to the heteroduplex substrate (21Lima W.F. Wu H. Nichols J. Prakash T.P. Ravikumar V. Crooke S.T. J. Biol. Chem. 2003; 278: 49860-49867Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar). The conserved tryptophan at position 43 is responsible for properly positioning the enzyme on the substrate for catalysis (21Lima W.F. Wu H. Nichols J. Prakash T.P. Ravikumar V. Crooke S.T. J. Biol. Chem. 2003; 278: 49860-49867Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar). Human RNase H1 exhibits a strong positional preference for cleavage, i.e. human RNase H1 cleaves the heteroduplex substrate between 7 and 12 nucleotides from the 3′-DNA/5′-RNA terminus (14Wu H. Lima W.F. Crooke S.T. J. Biol. Chem. 2001; 276: 23547-23553Abstract Full Text Full Text PDF PubMed Scopus (66) Google Scholar and 21Lima W.F. Wu H. Nichols J. Prakash T.P. Ravikumar V. Crooke S.T. J. Biol. Chem. 2003; 278: 49860-49867Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar). Based on site-directed mutagenesis of both human RNase H1 and the heteroduplex substrate, the RNA-binding domain was shown to be responsible for the observed positional preference for cleavage (21Lima W.F. Wu H. Nichols J. Prakash T.P. Ravikumar V. Crooke S.T. J. Biol. Chem. 2003; 278: 49860-49867Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar). The RNA-binding domain of human RNase H1 appears to bind to the 3′-DNA/5′-RNA pole of the heteroduplex substrate, with the catalytic site of the enzyme positioned slightly less than one helical turn from the RNA-binding domain (21Lima W.F. Wu H. Nichols J. Prakash T.P. Ravikumar V. Crooke S.T. J. Biol. Chem. 2003; 278: 49860-49867Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar). Substitution of either the terminal 3′-DNA with a single ribonucleotide or 5′-RNA with a 2′-methoxyethoxy deoxyribonucleotide was shown to cause a concomitant 3′-shift of the first 5′-cleavage site on the RNA, suggesting that altering duplex geometry interferes with proper positioning of the enzyme on the heteroduplex for cleavage (21Lima W.F. Wu H. Nichols J. Prakash T.P. Ravikumar V. Crooke S.T. J. Biol. Chem. 2003; 278: 49860-49867Abstract Full Text Full Text PDF PubMed Scopus (39) Google Scholar). Although the interaction between the RNA-binding domain and the heteroduplex substrate has been characterized, the mechanism by which the catalytic domain of RNase H1 recognizes the substrate has not been fully elucidated. Human RNase H1 is a nuclease that cleaves RNA exclusively in an RNA/DNA duplex via a double-strand DNase cleavage mechanism. Neither double-strand RNA nor double-strand DNA duplexes support RNase H1 activity (22Lima W.F. Crooke S.T. Biochemistry. 1997; 36: 390-398Crossref PubMed Scopus (115) Google Scholar, 23Wu H. Lima W.L. Crooke S.T. J. Biol. Chem. 1999; 274: 28270-28278Abstract Full Text Full Text PDF PubMed Scopus (144) Google Scholar). The observed structural differences between the RNA/DNA heteroduplex and double-strand RNA and double-strand DNA duplexes suggest a possible role for the helical geometry and sugar conformation of the DNA and RNA in the selective cleavage of the heteroduplex substrate by human RNase H1 (24Fedoroff O.Y. Salazar M. Reid B.R. J. Mol. Biol. 1993; 233: 509-523Crossref PubMed Scopus (234) Google Scholar, 25Egli M. Portman S. Usman N. Biochemistry. 1996; 35: 8489-8494Crossref PubMed Scopus (198) Google Scholar, 26Saenger W. Principles of Nucleic Acid Structure. Springer-Verlag New York Inc., New York1984Crossref Google Scholar). Specifically, the deoxyribonucleotides within double-strand DNA form a southern C-2′-endo sugar conformation, resulting in a B-form helical conformation, whereas ribonucleotides within double-strand RNA form a northern C-3′-endo pucker and an A-form helical geometry (26Saenger W. Principles of Nucleic Acid Structure. Springer-Verlag New York Inc., New York1984Crossref Google Scholar). In contrast, the deoxyribonucleotides of the RNA/DNA heteroduplex have been shown to adopt an eastern O-4′-endo sugar pucker, resulting in a helical conformation in which the RNA strand adopts A-form geometry and the DNA strand shares both the A- and B-form helical conformations. The conformational diversity observed for the DNA strand is likely a function of the intrinsic flexibility of the deoxyribonucleotide compared with RNA and may also be important for human RNase H1 activity. DNA also differs from RNA in that the furanose ring of deoxynucleotide is much more flexible, i.e. it exhibits a nearly symmetrical potential energy barrier for both southern and northern sugar conformations (26Saenger W. Principles of Nucleic Acid Structure. Springer-Verlag New York Inc., New York1984Crossref Google Scholar). Consistent with these observations, heteroduplexes containing 2′-arafluoro deoxyribonucleotides, which have been shown to exhibit a sugar conformation comparable with DNA when hybridized to RNA, have also been shown to support RNase H1 activity (27Denissov A.Y. Noronha A.M. Wilds C.J. Trempe J.-F. Pon R.T. Gehring K. Damha M. Nucleic Acids Res. 2001; 29: 4284-4293Crossref PubMed Scopus (50) Google Scholar). On the other hand, heteroduplexes consisting of RNA/2′-alkoxy-modified deoxyribonucleotides exhibiting a C-3′-endo sugar pucker and an A-form helical geometry when hybridized to RNA do not support human RNase H1 activity (22Lima W.F. Crooke S.T. Biochemistry. 1997; 36: 390-398Crossref PubMed Scopus (115) Google Scholar, 23Wu H. Lima W.L. Crooke S.T. J. Biol. Chem. 1999; 274: 28270-28278Abstract Full Text Full Text PDF PubMed Scopus (144) Google Scholar). We have previously shown that both E. coli and human RNases H1 bind A-form duplexes (e.g. RNA/RNA, 2′-methoxyethoxy/RNA, and 2′-methoxy/RNA) with comparable affinity to the RNA/DNA heteroduplex substrate but do not cleave the A-form duplexes (22Lima W.F. Crooke S.T. Biochemistry. 1997; 36: 390-398Crossref PubMed Scopus (115) Google Scholar, 23Wu H. Lima W.L. Crooke S.T. J. Biol. Chem. 1999; 274: 28270-28278Abstract Full Text Full Text PDF PubMed Scopus (144) Google Scholar). In this case, the size and position of the 2′-substituents of RNA and 2′-alkoxy nucleotides suggest possible steric interference with RNase H1, as the 2′-substituents are positioned within the minor groove of the heteroduplex, a region predicted to be the binding site for the enzyme (28Katayangi K. Okumura M. Morikawa K. Proteins Struct. Funct. Genet. 1993; 17: 337-346Crossref PubMed Scopus (141) Google Scholar). Alternatively, the sugar conformation and flexibility map play a decisive role in RNase H1 activity. In this study, we have determined the structure/activity relationships for the interaction between the catalytic domain of human RNase H1 and the RNA/DNA heteroduplex substrate by systematically evaluating the influence of sugar conformation, bulk in the minor groove, and flexibility in the catalytic area on enzyme efficiency. Modified nucleotides were introduced into the oligodeoxyribonucleotides at the human RNase H1 preferred cleavage sites on the heteroduplex and consisted of the DNA-like southern C-2′-endo, RNA-like northern C-3′-endo, and eastern O-4′-endo biased sugars with and without 2′-substituents (see Fig. 1A). In addition, varying degrees of conformational flexibility were introduced into the heteroduplex substrate by incorporating modified deoxyribonucleotides that ∏-stack with the adjacent deoxyribonucleotides but do not form hydrogen bonds with the bases of the RNA strand, abasic deoxynucleotides, hydrocarbon intranucleotide linkers, and the ganciclovir-modified deoxyribonucleotide (see Fig. 1B). The initial cleavage rates (V0) and the site-specific cleavage rates of the modified heteroduplexes were compared with those of the wild-type RNA/DNA heteroduplex. Preparation of Human RNase H1—Human RNase H1 containing an N-terminal His tag was expressed and purified as described previously (29Lima W.F. Wu H. Crooke S.T. Methods Enzymol. 2001; 341: 430-439Crossref PubMed Scopus (39) Google Scholar). Briefly, the plasmids were transfected into E. coli BL21(DE3) (Novagen). The bacteria was grown in Terrific Broth medium (Bio 101, Inc.) at 37 °C and harvested at A600 = 1.2. The cells were induced with 1 mm isopropyl-β-d-thiogalactopyranoside at 37 °C for 2 h. The cells were lysed in 6 m guanidine hydrochloride, 100 mm sodium phosphate, and 10 mm Tris (pH 8.0) for 16–20hat24 °C. The recombinant proteins were incubated for 1 h with nickel-nitrilotriacetic acid super flow beads (1 ml/50 ml of lysate; QIAGEN Inc.). The nickel-nitrilotriacetic acid medium was packed into a fast protein liquid chromatography column, and the RNase H1 proteins were partially purified at a flow rate of 5 ml/min with sequential gradients (buffer A = 100 mm sodium phosphate, 10 mm Tris-HCl, and 8 m urea (pH 6.3); buffer B = 100 mm sodium phosphate, 10 mm Tris-HCl, and 2 m urea (pH 6.3); and buffer C = 100 mm sodium phosphate, 10 mm Tris-HCl, 2 m urea, and 100 mm EDTA (pH 7.0)). The eluent was further purified by ion-exchange fast protein liquid chromatography on a Mono S column at a flow rate of 1 ml/min (buffer A = 20 mm sodium phosphate, 2 m urea, and 200 mm NaCl (pH 7.0) and buffer B = 20 mm sodium phosphate, 2 m urea, and 2 m NaCl (pH 7.0)). Fractions containing RNase H1 were pooled and concentrated. The concentrated protein was purified by reverse phase fast protein liquid chromatography on a Resourse RPC column at a flow rate of 1 ml/min (buffer A = 2% acetonitrile in distilled H2O and 0.065% trifluoroacetic acid and buffer B = 80% acetonitrile in distilled H2O and 0.05% trifluoroacetic acid). Fractions were lyophilized, resuspended in distilled H2O, and analyzed by SDS-PAGE. Synthesis of Oligonucleotides—The oligoribonucleotides were synthesized on a PE-ABI 380B synthesizer using 5′-O-silyl-2′-O-bis(2-acetoxyethoxy)methyl ribonucleoside phosphoramidites and procedures described elsewhere (30Scaringe S.A. Wincott F.E. Caruthers M.H. J. Am. Chem. Soc. 1998; 120: 11820-11821Crossref Scopus (217) Google Scholar). The oligoribonucleotides were purified by reverse-phase HPLC. The oligodeoxyribonucleotides were synthesized on a PE-ABI 380B automated DNA synthesizer using standard phosphoramidite chemistry. The oligodeoxyribonucleotides were purified by precipitation two times out of 0.5 m NaCl with 2.5 volumes of ethyl alcohol. Synthesis of Modified Oligonucleotides—1,4-Anhydro-5-O-(4,4′-dimethoxytrityl)-2-deoxy-d-erythro-pentenol-3-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite, 5′-O-(4,4′-dimethoxytrityl)-3-(4-methylbenzoyl)-2-thiothymidine-3′-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite, 5′-O-(4,4′-dimethoxytrityl)-3′-deoxypseuodouridine-3′-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite, 1′,2′-dideoxy-1′-(2,4-difluorotoluyl)-5′-O-(4,4′-dimethoxytrityl)-β-d-ribofuranose-3′-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite, and 5′-O-(4,4′-dimethoxytrityl)-3-(4-methylbenzoyl)-2′-uridine-3′-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite were procured from Glen Research Inc. 1-[2-Deoxy-2-fluoro-5-O-(4,4′-dimethoxytrityl)]-3-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite-β-d-[arabinofuranosyl]thymine, 5′-O-(4,4′-dimethoxytrityl)-2′-deoxy-2′-fluorothymidine-3′-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite, 2-fluoro-6-methylbenzoimidazole deoxyribonucleotide, 4-methylbenzoimidazole deoxyribonucleotide, hydrocarbon linkers, and 5′-O-(4,4′-dimethoxytrityl)-2′-S-methyl-2′-thio-5-methyluridine-3′-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite were synthesized as described (31Kawasaki A.M. Casper M.D. Freier S.M. Lesnik E.A. Zounes M.C. Cummins L.L. Gonzalez C. Cook P.D. J. Med. Chem. 1993; 36: 831-841Crossref PubMed Scopus (373) Google Scholar, 32Yoneda N. Tetrahedron. 1991; 47: 5329-5365Crossref Scopus (133) Google Scholar, 33Ikeda H. Fernandez R. Wilk A. Barchi Jr., J.J. Huang X. Marquez V.E. Nucleic Acids Res. 1998; 26: 2237-2244Crossref PubMed Scopus (81) Google Scholar). The nucleoside 3′-β-C-methylthymidine was synthesized from 1,2-O-isopropylidene-d-xylofuranose and converted to the 5′-O-(4,4′-dimethoxytrityl)-3′-O-(2-cyanoethyldiisopropylamino)phosphoramidite as described previously (34Wilds C.J. Damha M.J. Nucleic Acids Res. 2000; 28: 3625-3635Crossref PubMed Google Scholar, 52Fraser A. Wheeler P. Cook P.D. Sanghvi Y.S. J. Heterocycl. Chem. 1993; 30: 212-224Crossref Scopus (25) Google Scholar). The 4′-α-C-methylthymidine nucleoside was synthesized in 12 steps, 3J. Xia, S. S. Carroll, D. B. Olsen, M. MacCoss, P. D. Cook, A. B. Eldrup, F. Bennett, and B. Bhat, manuscript in preparation. starting from commercially available 1,2,5,6-di-O-isopropylidene-α-d-glucofuranose (Pfanstiehl Laboratories Inc., Waukegan, IL). The alternative synthesis of this nucleoside has been reported (35Sproat B.S. Gait M.J. Gait M.J. Oligonucleotide Synthesis: A Practical Approach. IRL Press, Washington, D. C.1985: 83-115Google Scholar, 36Schmit C. Bevierre M.-O. De Mesmaeker A. Altmann K.-H. Bioorg. Med. Chem. Lett. 1994; 4: 1969-1974Crossref Scopus (48) Google Scholar). It was converted to 5′-O-(4,4′-dimethoxytrityl)-4′-α-C-methylthymidine-3′-O-[(2-cyanoethyl)-N,N-diisopropyl]phosphoramidite following the procedure described for similar compounds (37Waga T. Ohurui H. Meguro H. Nucleosides Nucleotides. 1996; 15: 287-304Crossref Scopus (55) Google Scholar, 38Detmer I. Summerer D. Marx A. Eur. J. Org. Chem. 2003; 10: 1837-1846Crossref Scopus (21) Google Scholar). Standard phosphoramidites and solid supports were used for incorporation of A, T, G, and C residues. A 0.1 m solution of each amidite in anhydrous acetonitrile was used for the synthesis of modified oligonucleotides. The oligonucleotides were synthesized on functionalized Controlled-Pore Glass on an automated solid-phase DNA synthesizer with the final dimethoxytrityl group retained at 5′-end. For incorporation of modified amidites, 6 eq of phosphoramidite solutions were delivered in two portions, each followed by a 3-min coupling wait time. All other steps in the protocol supplied by the manufacturer were used without modification. Oxidation of the internucleotide phosphite to the phosphate was carried out using a 0.1 m solution of iodine in 20:1 (v/v) pyridine/water with a 10-min oxidation wait time. The coupling efficiencies were >97%. To deprotect oligonucleotides containing 2′-deoxy-2′-fluorothymidine and 2′-deoxy-2′-fluoroarabinofuranosylthymine, the solid supports bearing the oligonucleotides were suspended in aqueous ammonia (28–30 weight %)/ethanol (3:1; 3 ml for 2-μmol scale synthesis) and heated at 55 °C for 6 h. For all other modified oligonucleotides after completion of the synthesis, the solid supports bearing the oligonucleotides were suspended in aqueous ammonium hydroxide (28–30 weight %; 2 ml for 2-μmol scale synthesis) and kept at room temperature for 2 h. The solid support was filtered, and the filtrate was heated at 55 °C for 6 h to complete the removal of all protecting groups. Crude oligonucleotides were purified on a Waters HPLC C4 7.8 × 300-mm column (buffer A = 100 mm ammonium acetate (pH 6.5–7); buffer B = acetonitrile; 5–60% of buffer B in 55 min) at a flow rate of 2.5 ml/min (λ260 nm). Detritylation was achieved by adjusting the pH of the solution to 3.8 with acetic acid and by keeping at room temperature until complete removal of the trityl group, as monitored by HPLC analysis. The oligonucleotides were then desalted by HPLC to yield modified oligonucleotides in 30–40% isolated yield calculated based on the loading of the 3′-base onto the solid support (39Kanazaki M. Ueno Y. Shuto S. Matsuda A. J. Am. Chem. Soc. 2000; 122: 2422-2432Crossref Scopus (64) Google Scholar). The oligonucleotides were characterized by electrospray mass spectroscopy, and their purity was assessed by HPLC and capillary gel electrophoresis. The purity of the oligonucleotides was >90%. Oligonucleotides with abasic sites were conveniently generated by the use of uracil-DNA glycosylase (40Duncan B.K. Chambers J.A. Gene (Amst.). 1984; 28: 211-219Crossref PubMed Scopus (18) Google Scholar). Oligonucleotides containing deoxyuridine residues were synthesized as described above. The HPLC-purified oligonucleotides (0.32 mg) were dissolved in uracil-DNA glycosylase (149 μl; 1 unit in 1 μl dissolved in 30 mm HEPES-KOH (pH 7.5), 150 mm NaCl, 1 mm EDTA, 1 mm dithiothreitol, 0.05% Tween 20, and 50% glycerol) and incubated at 37 °C for 4 h. The reaction was terminated by filtering the enzyme using a low binding membrane filter (0.22 μm; Millipore Corp., Bedford, MA). The release of uracil was observed upon HPLC analysis of the reaction mixture using a Waters C4 3.9 × 300-mm column (Delta Pack, 15 μm, 300 Å) (buffer A = 100 mm ammonium acetate; buffer B = acetonitrile; 0–25% buffer B in 55 min) at a flow rate of 1.5 ml/min (λ260 nm) and co-injection of the authentic sample. The oligonucleotides were purified by HPLC as described above. The purity (>90%) of the oligonucleotides was assessed by HPLC analysis. Preparation of 32P-Labeled Substrate—The RNA substrate was 5′-end-labeled with 32P using 20 units of T4 polynucleotide kinase (Promega Corp.), 120 pmol (7000 Ci/mmol) of [γ-32P]ATP (ICN), 40 pmol of RNA, 70 mm Tris (pH 7.6), 10 mm MgCl2, and 50 mm dithiothreitol. The kinase reaction was incubated at 37 °C for 30 min. The labeled oligoribonucleotide was purified by electrophoresis on a 12% denaturing polyacrylamide gel (41Sambrook J. Fritsch E.F. Maniatis T. Molecular Cloning: A Laboratory Manual. 2nd Ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY1989Google Scholar). The specific activity of the labeled oligonucleotide is ∼3000–8000 cpm/fmol. Preparation of the Heteroduplex—The heteroduplex substrate was prepared in 100 μl containing 100–1000 nm unlabeled oligoribonucleotide, 105 cpm 32P-labeled oligoribonucleotide, 2-fold excess complementary oligodeoxyribonucleotide, and hybridization buffer (20 mm Tris (pH 7.5) and 20 mm KCl). Reactions were heated at 90 °C for 5 min and cooled to 37 °C, and 60 units of Prime RNase Inhibitor (5 Prime → 3 Prime, Inc., Boulder, CO) and MgCl2 at a final concentration of 1 mm were added. Hybridization reactions were incubated for 2–16hat37 °C, and 1 mm tris(2-carboxyethyl) phosphate was added. Multiple Turnover Kinetics—The human RNase H1 proteins were incubated with dilution buffer (50 mm Tris, 50 mm NaCl, and 100 μm tris(2-carboxyethyl) phosphate (pH 7.5)) for 1 h at 24 °C. The heteroduplex substrate was digested with 0.4 ng of enzyme at 37 °C. A 10-μl aliquot of the cleavage reaction was removed at time points ranging from 2 to 120 min and quenched by adding 5 μl of stop solution (8 m urea and 120 mm EDTA). The aliquots were heated at 90 °C for 2 min and resolved on a 12% denaturing polyacrylamide gel, and the substrate and product bands were quantitated on a Amersham Biosciences PhosphorImager. The concentration of the converted product was plotted as a function of time. The initial cleavage rate (V0) was obtained from the slope (mo