Relationship between the Oligomeric Status of HIV-1 Integrase on DNA and Enzymatic Activity
2006; Elsevier BV; Volume: 281; Issue: 32 Linguagem: Inglês
10.1074/jbc.m602198200
ISSN1083-351X
AutoresElvire Guiot, Kévin Carayon, Olivier Delelis, Françoise Simon, Patrick Tauc, E. M. Zubin, Marina Gottikh, Jean‐François Mouscadet, Jean‐Claude Brochon, Eric Deprez,
Tópico(s)HIV Research and Treatment
ResumoThe 3′-processing of the extremities of viral DNA is the first of two reactions catalyzed by HIV-1 integrase (IN). High order IN multimers (tetramers) are required for complete integration, but it remains unclear which oligomer is responsible for the 3′-processing reaction. Moreover, IN tends to aggregate, and it is unknown whether the polymerization or aggregation of this enzyme on DNA is detrimental or beneficial for activity. We have developed a fluorescence assay based on anisotropy for monitoring release of the terminal dinucleotide product in real-time. Because the initial anisotropy value obtained after DNA binding and before catalysis depends on the fractional saturation of DNA sites and the size of IN·DNA complexes, this approach can be used to study the relationship between activity and binding/multimerization parameters in the same assay. By increasing the IN:DNA ratio, we found that the anisotropy increased but the 3′-processing activity displayed a characteristic bell-shaped behavior. The anisotropy values obtained in the first phase were predictive of subsequent activity and accounted for the number of complexes. Interestingly, activity peaked and then decreased in the second phase, whereas anisotropy continued to increase. Time-resolved fluorescence anisotropy studies showed that the most competent form for catalysis corresponds to a dimer bound to one viral DNA end, whereas higher order complexes such as aggregates predominate during the second phase when activity drops off. We conclude that a single IN dimer at each extremity of viral DNA molecules is required for 3′-processing, with a dimer of dimers responsible for the subsequent full integration. The 3′-processing of the extremities of viral DNA is the first of two reactions catalyzed by HIV-1 integrase (IN). High order IN multimers (tetramers) are required for complete integration, but it remains unclear which oligomer is responsible for the 3′-processing reaction. Moreover, IN tends to aggregate, and it is unknown whether the polymerization or aggregation of this enzyme on DNA is detrimental or beneficial for activity. We have developed a fluorescence assay based on anisotropy for monitoring release of the terminal dinucleotide product in real-time. Because the initial anisotropy value obtained after DNA binding and before catalysis depends on the fractional saturation of DNA sites and the size of IN·DNA complexes, this approach can be used to study the relationship between activity and binding/multimerization parameters in the same assay. By increasing the IN:DNA ratio, we found that the anisotropy increased but the 3′-processing activity displayed a characteristic bell-shaped behavior. The anisotropy values obtained in the first phase were predictive of subsequent activity and accounted for the number of complexes. Interestingly, activity peaked and then decreased in the second phase, whereas anisotropy continued to increase. Time-resolved fluorescence anisotropy studies showed that the most competent form for catalysis corresponds to a dimer bound to one viral DNA end, whereas higher order complexes such as aggregates predominate during the second phase when activity drops off. We conclude that a single IN dimer at each extremity of viral DNA molecules is required for 3′-processing, with a dimer of dimers responsible for the subsequent full integration. The integration of a DNA copy of the HIV-1 2The abbreviations used are: HIV, human immunodeficiency virus; IN, integrase; LTR, long terminal repeat; ODN, oligodeoxynucleotide; ds, double-stranded; ss, single-stranded; r, steady-state fluorescence anisotropy; TFA, time-resolved fluorescence anisotropy; TAMRA, carboxytetramethylrhodamine; wt, wild-type.2The abbreviations used are: HIV, human immunodeficiency virus; IN, integrase; LTR, long terminal repeat; ODN, oligodeoxynucleotide; ds, double-stranded; ss, single-stranded; r, steady-state fluorescence anisotropy; TFA, time-resolved fluorescence anisotropy; TAMRA, carboxytetramethylrhodamine; wt, wild-type. genome into the host genome is a crucial step in the life cycle of the retrovirus. Integrase (IN) is responsible for the two consecutive reactions that constitute the overall integration process. The first of these two reactions is 3′-processing, which involves cleavage of the 3′-terminal GT dinucleotide at each extremity of the viral DNA. The hydroxyl groups of newly recessed 3′-ends are then used in the second reaction, strand transfer, for the covalent joining of viral and target DNAs, resulting in full-site integration. IN is sufficient for catalysis of the 3′-processing reaction in vitro, using short-length oligodeoxynucleotides (ODNs) that mimic one viral long terminal repeat (LTR) in the presence of the metallic cofactor Mg2+. This reaction generates two products: the viral DNA containing the recessed extremity and the GT dinucleotide. One of the two products, the processed viral DNA, as well as the target DNA serve as substrates for the subsequent joining reaction. IN belongs to the superfamily of polynucleotidyl transferases. Its catalytic core domain contains a triad of acidic residues constituting the D,D-35-E motif, which is strictly required for catalysis. The catalytic core establishes specific contacts with the viral DNA and, together with the C-terminal domain, is involved in DNA binding (1Lutzke R.A. Vink C. Plasterk R.H. Nucleic Acids Res. 1994; 22: 4125-4131Crossref PubMed Scopus (172) Google Scholar, 2Engelman A. Hickman A.B. Craigie R. J. Virol. 1994; 68: 5911-5917Crossref PubMed Google Scholar, 3Jenkins T.M. Esposito D. Engelman A. Craigie R. EMBO J. 1997; 16: 6849-6859Crossref PubMed Scopus (215) Google Scholar, 4Esposito D. Craigie R. EMBO J. 1998; 17: 5832-5843Crossref PubMed Scopus (260) Google Scholar). The 3′-processing reaction is highly specific, and the terminal 13 bp of the LTR play a key role for Mg2+-dependent 3′-processing in terms of reaction specificity (4Esposito D. Craigie R. EMBO J. 1998; 17: 5832-5843Crossref PubMed Scopus (260) Google Scholar, 5Agapkina J. Smolov M. Barbe S. Zubin E. Zatsepin T. Deprez E. Le Bret M. Mouscadet J.F. Gottikh M.V. J. Biol. Chem. 2006; 281: 11530-11540Abstract Full Text Full Text PDF PubMed Scopus (37) Google Scholar). In particular, the CA sequence preceding the GT dinucleotide cleaved by IN is strictly required. Reaction specificity seems to depend on the catalytic step, because no significant difference in affinity is observed in vitro for different DNA sequences (2Engelman A. Hickman A.B. Craigie R. J. Virol. 1994; 68: 5911-5917Crossref PubMed Google Scholar, 3Jenkins T.M. Esposito D. Engelman A. Craigie R. EMBO J. 1997; 16: 6849-6859Crossref PubMed Scopus (215) Google Scholar, 5Agapkina J. Smolov M. Barbe S. Zubin E. Zatsepin T. Deprez E. Le Bret M. Mouscadet J.F. Gottikh M.V. J. Biol. Chem. 2006; 281: 11530-11540Abstract Full Text Full Text PDF PubMed Scopus (37) Google Scholar, 6Mazumder A. Neamati N. Pilon A.A. Sunder S. Pommier Y. J. Biol. Chem. 1996; 271: 27330-27338Abstract Full Text Full Text PDF PubMed Scopus (58) Google Scholar). The catalytic mechanism of IN has been extensively studied, but the structural determinants of IN activity remain unclear, and controversy remains concerning the multimeric status of active IN. Single- and double-domain crystallographic structures have been produced for IN, alone or complexed with the IN-binding domain of the cellular partner lens epithelium-derived growth factor, and all these structures display a conserved dimeric structure of the catalytic core (7Wang J.Y. Ling H. Yang W. Craigie R. EMBO J. 2001; 20: 7333-7343Crossref PubMed Scopus (311) Google Scholar, 8Maignan S. Guilloteau J.P. Zhou-Liu Q. Clement-Mella C. Mikol V. J. Mol. Biol. 1998; 282: 359-368Crossref PubMed Scopus (265) Google Scholar, 9Chen J.C. Krucinski J. Miercke L.J. Finer-Moore J.S. Tang A.H. Leavitt A.D. Stroud R.M. Proc. Natl. Acad. Sci. U. S. A. 2000; 97: 8233-8238Crossref PubMed Scopus (380) Google Scholar, 10Goldgur Y. Dyda F. Hickman A.B. Jenkins T.M. Craigie R. Davies D.R. Proc. Natl. Acad. Sci. U. S. A. 1998; 95: 9150-9154Crossref PubMed Scopus (366) Google Scholar, 11Cherepanov P. Ambrosio A.L. Rahman S. Ellenberger T. Engelman A. Proc. Natl. Acad. Sci. U. S. A. 2005; 102: 17308-17313Crossref PubMed Scopus (347) Google Scholar). Based on topological considerations concerning the distance separating the active sites of the two protomers, which is more than double the distance separating the two joining sites on the target DNA (separated by 5 bp), a multimeric state of an order higher than dimers, tetramers at least, is required for complete integration. However, it remains unclear whether such a higher order multimeric state is also required for 3′-processing activity and how this activity is modulated by the self-association properties of IN. Moreover, most enzymatic studies are carried out with an excess of IN over DNA substrate. Taking into account the low solubility of IN, we investigated whether high order multimers or aggregated forms of IN favored by high protein concentrations in solution were detrimental or beneficial for activity. We developed an assay for monitoring DNA binding and subsequent 3′-processing in the same sample. This assay makes possible the separation of binding and catalytic parameters and the study of real-time kinetics. It is based on steady-state fluorescence anisotropy (r), which is sensitive to rotational diffusion, and thus suitable for studies aiming to identify structural modifications leading to a significant change in molecular size (12Deprez E. Barbe S. Kolaski M. Leh H. Zouhiri F. Auclair C. Brochon J.C. Le Bret M. Mouscadet J.F. Mol. Pharmacol. 2004; 65: 85-98Crossref PubMed Scopus (86) Google Scholar, 13Yang J. Xi J. Zhuang Z. Benkovic S.J. J. Biol. Chem. 2005; 280: 25416-25423Abstract Full Text Full Text PDF PubMed Scopus (28) Google Scholar, 14Weinberg R.L. Veprintsev D.B. Fersht A.R. J. Mol. Biol. 2004; 341: 1145-1159Crossref PubMed Scopus (191) Google Scholar, 15Grillo A.O. Brown M.P. Royer C.A. J. Mol. Biol. 1999; 287: 539-554Crossref PubMed Scopus (40) Google Scholar). Using a fluorescent probe covalently linked to the GT dinucleotide, this makes it possible to follow DNA binding and dinucleotide release, because both steps strongly influence molecular size of the fluorescent moiety. The anisotropy-based technique is also highly suitable for studies of the relationship between the overall size of IN·DNA complexes and activity. We found that, for low IN:DNA ratios, the r values obtained after DNA binding and before catalysis were fully predictive of subsequent IN activity, according to the fractional saturation function. For high IN:DNA ratios, anisotropy continued to increase, but 3′-processing activity decreased. Because r depends on both fractional saturation and the molecular size of complexes at saturation of DNA sites, our results show that high order multimers or aggregated states of IN are detrimental to 3′-processing activity. Activity levels were highest for non-aggregative smaller species. A more precise characterization of catalytically competent complexes by time-resolved fluorescence anisotropy (TFA) (16Deprez E. Tauc P. Leh H. Mouscadet J.F. Auclair C. Hawkins M.E. Brochon J.C. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 10090-10095Crossref PubMed Scopus (82) Google Scholar, 17Deprez E. Tauc P. Leh H. Mouscadet J.F. Auclair C. Brochon J.C. Biochemistry. 2000; 39: 9275-9284Crossref PubMed Scopus (128) Google Scholar, 18Ciubotaru M. Ptaszek L.M. Baker G.A. Baker S.N. Bright F.V. Schatz D.G. J. Biol. Chem. 2003; 278: 5584-5596Abstract Full Text Full Text PDF PubMed Scopus (25) Google Scholar, 19Perez-Howard G.M. Weil P.A. Beechem J.M. Biochemistry. 1995; 34: 8005-8017Crossref PubMed Scopus (131) Google Scholar, 20Bloom L.B. Turner J. Kelman Z. Beechem J.M. O'Donnell M. Goodman M.F. J. Biol. Chem. 1996; 271: 30699-30708Abstract Full Text Full Text PDF PubMed Scopus (53) Google Scholar), confirmed steady-state data and led to the identification of dimeric forms as the most active forms for 3′-processing. Our results also highlight that only the IN:DNA ratio but not the IN concentration per se determines the aggregation properties and thus the activity of IN and that DNA binding stimulates the self-organization of IN to give a catalytically competent non-aggregative form. Oligonucleotides and Nomenclature—Unlabeled and fluorescein-labeled single-stranded (ss) ODNs were purchased from Eurogentec (Liege, Belgium) (except for the 2′-aminouridine-containing ODN) and purified by electrophoresis in acrylamide gels. The specific HIV sequence was 5′-GTGTGGAAAATCTCTAGCAGT-3′ (the bases removed by IN are underlined). This sequence was denoted a, and the complementary nonprocessed strand was denoted b. Fluorescein (F) was attached at the 5′- or the 3′-end of strand a or b, via a 6-carbon linker. As an example, the specific double-stranded (ds) ODN used to monitor activity was called HIV-a3F, indicating that fluorescein was attached to the 3′-end of strand a. HIV-GTGT-a3F was identical except that the four terminal bases at the 3′-end of strand a were GTGT rather than the canonical sequence CAGT. The nonspecific sequence, NS-TTCC, was 5′-ACCTATGCGCCGCTAGATTCC-3′ (strand a). Two other nonspecific ODNs with different 3′-ends of strand a were derived: NS-CACC and NS-CAGT. ds ODNs were obtained by mixing equimolar amounts of complementary strands in 20 mm Tris-HCl (pH 7.2), 100 mm NaCl. The mixture was heated to 85 °C for 5 min, and annealing was allowed by slow cooling to 25 °C. The uridine-containing ODN (corresponding to strand a of the DNA substrate HIV-aUF) contains a fluorescein attached to the 2′-amino group of the 3′-terminal 2′-aminouridine. It was synthesized as follows: the solid support was prepared as previously described (21Kuznetsova L.G. Romanova E.A. Volkov E.M. Tashlitskii V.N. Oretskaia T.S. Krynetskaia N.F. Shabarova Z.A. Bioorg. Khim. 1993; 19: 455-466PubMed Google Scholar). Briefly, 2′-deoxy-5′-O-4,4′-dimethoxytrityl-2′-O-trifluoracetamidouridine (0.2 mmol) was co-evaporated with pyridine (3 × 5 ml) and dissolved in dry pyridine (2 ml). Succinylated long-chain alkylamine-controlled pore glass (500 Å) (200 mg), 2,4,6-triisopropylbenzenesulfonyl chloride (0.6 mmol), and 1-methylimidazole (1.2 mmol) were added, and the mixture was incubated for 3 h at 25°C. The support was then filtered and washed successively with pyridine, CH2Cl2, and ether. Starting from 2′-deoxy-5′-O-4,4′-dimethoxytrityl-2′-O-trifluoracetamidouridine-derivatized long-chain alkylamine-controlled pore glass (500 Å), a 21-mer ODN was assembled on an ABI394B DNA synthesizer, by the phosphoramidite method, according to the manufacturer's recommendation. Protected 2′-O-deoxyribonucleoside phosphoramidites and S-ethylthiotetrazole were purchased from Glen Research. Ammonia was used for cleavage from the support and deprotection overnight at 55 °C. The reaction mixture was then analyzed by reverse-phase high-performance liquid chromatography in ion-pair mode. For fluorescein labeling, 1 mg of fluorescein isothiocyanate dissolved in 20 μl of dimethylformamide was added to a solution of 40 nmol of 2′-amino ODN in 130 μl of sodium carbonate-bicarbonate (1 m, pH 9):water (v/v, 5:8) (22Antsypovich S.I. Volkov E.M. Oretskaia T.S. Romanova E.A. Tashlitskii V.N. Blumenfeld M. Shabarova Z.A. Bioorg. Khim. 1995; 21: 774-780PubMed Google Scholar, 23Agrawal S. Christodoulou C. Gait M.J. Nucleic Acids Res. 1986; 14: 6227-6245Crossref PubMed Scopus (197) Google Scholar). The mixture was incubated at 25 °C in the dark for 18 h and then loaded onto an NAP5-Sephadex G-25 column (Amersham Biosciences) pre-equilibrated in water. The ODN was eluted with water and analyzed by reversed-phase high-performance liquid chromatography in ion-pair mode. IN Purification and Standard Analysis of the 3′-Processing Reaction—HIV-1 IN (32 kDa) was purified under native conditions, as previously described (24Leh H. Brodin P. Bischerour J. Deprez E. Tauc P. Brochon J.C. LeCam E. Coulaud D. Auclair C. Mouscadet J.F. Biochemistry. 2000; 39: 9285-9294Crossref PubMed Scopus (120) Google Scholar). Standard IN assay, using 32P-labeled ODN and gel electrophoresis, was carried out as previously described (24Leh H. Brodin P. Bischerour J. Deprez E. Tauc P. Brochon J.C. LeCam E. Coulaud D. Auclair C. Mouscadet J.F. Biochemistry. 2000; 39: 9285-9294Crossref PubMed Scopus (120) Google Scholar). Gels were analyzed on a STORM 840™ PhosphorImager (Amersham Biosciences) and quantified with ImageQuaNT™ 4.1 software. The 3′-processing activity was calculated as follows: activity (%) = 19-mer/(21-mer + 19-mer) × 100. Steady-state Fluorescence Anisotropy Assay—Steady-state anisotropy values were recorded on a Beacon 2000 instrument (PanVera, Madison, WI), in a cell thermostatically held at 25 or 37 °C, for the DNA-binding step and 3′-processing reaction, respectively. Unless otherwise stated, we studied the formation of IN·DNA complexes by incubating fluorescein-labeled ds ODNs with IN in 20 mm Tris (pH 7.2), 1 mm dithiothreitol, 20 mm NaCl, 5 mm MgCl2 (the sample volume was 200 μl). The fractional saturation function (Y) was calculated as follows,Y=r-rODNrmax-rODN×100(Eq. 1) where rmax and rODN are the anisotropies of IN-bound and free ODN, respectively (no significant concomitant change in fluorescence intensity was observed). After the DNA-binding step, the temperature was raised from 25 to 37 °C for monitoring of the catalytic process. The 3′-processing activity was assessed by quantifying the decrease in r. Two independent methods were used for quantification as follows. (i) In fixed-time experiments, the reaction was stopped by adding SDS (0.25% final), disrupting all the IN·DNA complexes in the sample. In such experiments, the solution contained two fluorescent species: the non-processed ODN and the fluorescein-labeled dinucleotide released by the cleavage reaction. The fraction of dinucleotides released (Fdinu = [GT]/[DNA]total) is given by Equation 2,Fdinu=rNP-rrNP-rdinu(Eq. 2) where rNP and rdinu are the anisotropy values for pure solutions of non-processed ds ODN and dinucleotide, respectively (fluorescence did not change significantly during the reaction). We used the 5′-GT-3′F dinucleotide (Eurogentec) to determine rdinu. (ii) In real-time conditions, an additional fluorescent population corresponding to IN complexed with the unprocessed ds ODN, is present in the sample. In this case, Fdinu was calculated as follows,Fdinu=rt=0-rrmax-rdinu(Eq. 3) where rmax is the characteristic r value obtained for optimal activity, and rt = 0 is the r value obtained at the end of the DNA-binding step (before the start of the reaction). The 3′-processing activity obtained with Equations 2 and 3 is referred to as ActivitySDS and Activityreal-time, respectively. Activityreal-time was not used if rt = 0 was higher than 0.22 (aggregation of IN on DNA not negligible). We analyzed single-turnover kinetics using the Equations 4 and 5,ln(1-Fdinu)=-kobs×t(Eq. 4) withkobs=kchemistry/Kd,app/IN0+1(Eq. 5) where kchemistry is the single-turnover rate constant, and Kd,app is the apparent Kd (25Smolov M. Gottikh M. Tashlitskii V. Korolev S. Demidyuk I. Brochon J.C. Mouscadet J.F. Deprez E. FEBS J. 2006; 273: 1137-1151Crossref PubMed Scopus (39) Google Scholar). Reactions were conducted in the presence of Mg2+ (not Mn2+) to limit nonspecific hydrolysis products (cleavages at positions −3, −4, etc.), because anisotropy cannot discriminate between these small products (which are minor products with Mg2+ but not with Mn2+) and the specific GT product. The r values for the TAMRA-labeled DNA substrate (λex = 562 nm and λem = 582 nm) were recorded on a Cary Eclipse spectrofluorometer (Varian, Mulgrave, Australia) in polarization mode and were compared with values obtained with the same instrument and HIV-a3F as the substrate. Kd,app as a function of ODN length were determined by competition experiments. Fluorescein-labeled ds ODN HIV-a5F (4 nm) was preincubated with various concentrations of unlabeled ss or ds HIV ODN (from 10- to 45-mers) in 20 mm Tris buffer (pH 7.2) supplemented with 1 mm dithiothreitol, 20 mm NaCl, and 5 mm MgCl2. IN (150 nm final concentration) was then added, and r values were recorded. Δr (= r–rODN) was plotted against competitor concentration to determine Kd,app (concentration of competitor decreasing the initial Δr value by 50%). Time-resolved Fluorescence Experiments—Time-resolved fluorescence parameters (lifetimes and correlation times) were obtained from the two polarized fluorescence decays I∥(t) and I∥(t), using the time-correlated single photon counting technique. The instrumentation setup was essentially similar to those previously described (16Deprez E. Tauc P. Leh H. Mouscadet J.F. Auclair C. Hawkins M.E. Brochon J.C. Proc. Natl. Acad. Sci. U. S. A. 2001; 98: 10090-10095Crossref PubMed Scopus (82) Google Scholar, 17Deprez E. Tauc P. Leh H. Mouscadet J.F. Auclair C. Brochon J.C. Biochemistry. 2000; 39: 9275-9284Crossref PubMed Scopus (128) Google Scholar), with modifications: the time scaling was 19.5 ps per channel and 4096 channels were used. The excitation light pulse source was a Ti:sapphire laser (Millennia-pumped Tsunami femtosecond laser, Spectra Physics) (repetition rate: 8 MHz) associated with a second harmonic generator tuned to 490 nm. The emission monochromator (ARC SpectraPro-150) was set to 530 nm (Δλ = 15 nm). The two polarized components were collected alternately over a period of 30 s (total count of I∥: 15,000,000). The reaction mixture contained 20 mm Tris, pH 7.2, 20 mm NaCl, 5 mm MgCl2, 1 mm dithiothreitol. The correlation time (θ) distributions of free ss or ds ODNs were obtained at ODN concentrations of 10 nm or 0.5 μm. All ODNs (from 10- to 45-mer) mimicked the U5-end of the HIV-1 DNA and were fluorescein-labeled at the 5′-end of strand a. IN·DNA complexes were analyzed using ds HIV-a5F and various IN:DNA ratios, from 40 to 400. We analyzed both decays, I∥(t) and I∥(t), by the maximum entropy method (26Brochon J.C. Methods Enzymol. 1994; 240: 262-311Crossref PubMed Scopus (254) Google Scholar). Fluorescence anisotropy decay is described by Equations 6 and 7,r(t)=∑i=1nρi×e-t/θi(Eq. 6) with∑i=1nρi=ρ0(Eq. 7) where θi is the individual rotational correlation time, and ρi is the associated amplitude. ρ0 was found to lie between 0.36 and 0.37. Normalization of θ for a given temperature was performed using,θ=ηV/kT(Eq. 8) where η is the viscosity, V is the volume of the rotating unit, k is the Boltzmann constant, and T is the temperature (K). Monitoring 3′-Processing Activity by Steady-state Fluorescence Anisotropy—Fluorescence anisotropy measurements are based on the principle of photoselective excitation of a fluorophore by a polarized light, providing information about rotational motions of the fluorophore or fluorescently labeled molecule between photon absorption and emission. Some events such as overall rotational diffusion or flexibility are major causes of light depolarization. High levels of anisotropy are generally associated with large molecules or complexes characterized by slow rotational diffusion or low flexibility level. In this study, we used an extrinsic fluorophore covalently linked to DNA to monitor the binding of IN to viral DNA substrate and the subsequent 3′-processing reaction, in the same assay. Both DNA binding and 3′-processing would be expected to have a significant effect on the anisotropy parameter, because each of these steps has a major effect on the molecular size of the fluorescent moiety. The principle underlying the anisotropy-based assay is summarized in Fig. 1. The binding of IN to a ds ODN mimicking one end of the viral DNA increases the steady-state anisotropy value (r), most likely by restricting flexibility, allowing the calculation of fractional saturation function. Following DNA binding, the same sample can be shifted to a permissive temperature for 3′-processing activity. When the fluorophore is linked to the 3′-terminal GT dinucleotide, release of the dinucleotide product should significantly decrease r. The fraction of dinucleotides released can be determined from the real-time decrease in r observed during the reaction (Δrreal-time, corresponding to the difference between IN·DNA complex and free dinucleotide) or after the disruption of all IN·DNA complexes by the addition of SDS (ΔrSDS, corresponding to the difference between unprocessed ds ODN and free dinucleotide). Unlike most standard 3′-processing assays, which are based on quantification of the processed strand (first reaction product), the anisotropy-based assay monitors the released dinucleotide (second reaction product). We tested 21-mer DNA substrates labeled with fluorescein at one end, in all possible combinations (5′- or 3′-extremity of the a or b strand), or labeled on the 2′-amino group of a 3′-terminal 2′-aminouridine (Fig. 2A). The free ds ODNs were characterized by r values of 0.060–0.130 at 25 °C (0.045–0.105 at 37 °C), depending on the location of fluorescein, whereas the fluorescein-labeled dinucleotide was characterized by r = 0.02. DNA binding on the addition of IN was monitored at 25 °C and led to a significant increase in r value (>0.2 in the experimental conditions of Fig. 2). Equilibrium was typically reached after 15 min, as previously reported (12Deprez E. Barbe S. Kolaski M. Leh H. Zouhiri F. Auclair C. Brochon J.C. Le Bret M. Mouscadet J.F. Mol. Pharmacol. 2004; 65: 85-98Crossref PubMed Scopus (86) Google Scholar, 27Pinskaya M. Romanova E. Volkov E. Deprez E. Leh H. Brochon J.C. Mouscadet J.F. Gottikh M. Biochemistry. 2004; 43: 8735-8743Crossref PubMed Scopus (28) Google Scholar). The processing reaction was started by shifting the sample to a temperature of 37 °C, and the initial r values obtained therefore correspond to rt = 0. In the presence of the divalent cationic cofactor Mg2+, all tested ODNs gave similar rt = 0 values (0.22), indicating that fluorescein position had no effect on DNA binding by IN (Fig. 2B). Experiments carried out in the absence of Mg2+ gave reproducibly larger r values (0.25). This result confirms that DNA-binding activity of IN is not strictly dependent on Mg2+, consistent with other studies (2Engelman A. Hickman A.B. Craigie R. J. Virol. 1994; 68: 5911-5917Crossref PubMed Google Scholar, 28Hazuda D.J. Wolfe A.L. Hastings J.C. Robbins H.L. Graham P.L. LaFemina R.L. Emini E.A. J. Biol. Chem. 1994; 269: 3999-4004Abstract Full Text PDF PubMed Google Scholar, 29Vink C. Lutzke R.A. Plasterk R.H. Nucleic Acids Res. 1994; 22: 4103-4110Crossref PubMed Scopus (65) Google Scholar). Nevertheless, the larger r value suggests that IN aggregation is favored by an absence of Mg2+, as previously reported (17Deprez E. Tauc P. Leh H. Mouscadet J.F. Auclair C. Brochon J.C. Biochemistry. 2000; 39: 9275-9284Crossref PubMed Scopus (128) Google Scholar). The time dependence of r and Δr under real-time conditions at 37 °C is shown in Fig. 2 (B and C): only the DNA substrates HIV-a3F and HIV-aUF, with the fluorescein directly attached to the small GT reaction product, displayed significant decreases in r or Δr, and these decreases were strictly related to the presence of the metallic cofactor. In contrast, HIV-a5F, HIV-b5F, and HIV-b3F displayed no significant decrease in r or Δr value, under either real-time (Fig. 2) or fixed-time conditions (data not shown) (3′-processing activity was normally detected by a gel-electrophoresis method for HIV-b5F and HIV-b3F; HIV-a5F was not tested). The result obtained with these three ODNs indicates that anisotropy is not sensitive enough to differentiate between the DNA substrate (ds ODN 21/21) and the first reaction product (ds ODN 19/21). Furthermore, under real-time conditions, this result suggests that the processed DNA product remains tightly bound to the enzyme after 3′-processing. This tight binding together with the slow catalytic step could be responsible for the observed single-turnover property of IN, even under conditions of excess DNA substrate (25Smolov M. Gottikh M. Tashlitskii V. Korolev S. Demidyuk I. Brochon J.C. Mouscadet J.F. Deprez E. FEBS J. 2006; 273: 1137-1151Crossref PubMed Scopus (39) Google Scholar). In contrast, the terminal dinucleotide is released normally from IN·DNA complexes after 3′-processing, and our results indicate that the decrease in r is related to the formation of the GT product. Thus, anisotropy is reliable for monitoring the 3′-processing reaction, but only if fluorescein is attached to the GT dinucleotide. The 3′-processing reaction is strongly sequence-dependent, and the endonucleolytic site includes the crucial conserved CA dinucleotide immediately preceding the GT dinucleotide. We assessed sequence specificity, using the anisotropy-based assay and the following ODNs: HIV-a3F corresponds to the wild-type (wt) sequence, whereas HIV-GTGT-a3F is a variant in which the 3′-terminal CAGT sequence is replaced by GTGT. We also tested three nonspecific sequences, NS-TTCC-a3F, NS-CACC-a3F, and NS-CAGT-a3F. We found that the DNA-binding step was not influenced by sequence. Indeed, the final r values obtained after DNA binding were similar for all ODNs, irrespective of sequence context, confirming that in vitro assays primarily reveal the nonspecific DNA binding mode of IN. In contrast, sequence had a major effect on Δr value in catalysis conditions (Fig. 3A). Only HIV-a3F gave a large decrease in Δr. This decrease was abolished by replacement of the 3′-terminal sequence CAGT by GTGT or the use of nonspecific sequences, even with the CAGT sequence at the 3′-end. These results confirm that the CA dinucleotide is strictly required but not sufficient for activity, particularly in the presence of Mg2+, which gives more stringent conditions than Mn2+ (
Referência(s)