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Is a Closing “GA Pair” a Rule for Stable Loop-Loop RNA Complexes?

2000; Elsevier BV; Volume: 275; Issue: 28 Linguagem: Inglês

10.1074/jbc.m002694200

1083-351X

Frédéric Ducongé, Carmelo Di Primo, Jean‐Jacques Toulmé,

RNA Research and Splicing

RNA hairpin aptamers specific for the trans-activation-responsive (TAR) RNA element of human immunodeficiency virus type 1 were identified by in vitro selection (Ducongé, F., and Toulmé, J. J. (1999) RNA5, 1605–1614). The high affinity sequences selected at physiological magnesium concentration (3 mm) were shown to form a loop-loop complex with the targeted TAR RNA. The stability of this complex depends on the aptamer loop closing "GA pair" as characterized by preliminary electrophoretic mobility shift assays. Thermal denaturation monitored by UV-absorption spectroscopy and binding kinetics determined by surface plasmon resonance show that the GA pair is crucial for the formation of the TAR-RNA aptamer complex. Both thermal denaturation and surface plasmon resonance experiments show that any other "pairs" leads to complexes whose stability decreases in the order AG > GG > GU > AA > GC > UA >> CA, CU. The binding kinetics indicate that stability is controlled by the off-rate rather than by the on-rate. Comparison with the complex formed with the TAR* hairpin, a rationally designed TAR RNA ligand (Chang, K. Y., and Tinoco, I. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 8705–8709), demonstrates that the GA pair is a key determinant which accounts for the 50-fold increased stability of the TAR-aptamer complex (K d= 2.0 nm) over the TAR-TAR* one (K d = 92.5 nm) at physiological concentration of magnesium. Replacement of the wild-type GC pair next to the loop of RNA I′ by a GA pair stabilizes the RNA I′-RNA II′ loop-loop complex derived from the one involved in the control of the ColE1 plasmid replication. Thus, the GA pair might be the preferred one for stable loop-loop interactions. RNA hairpin aptamers specific for the trans-activation-responsive (TAR) RNA element of human immunodeficiency virus type 1 were identified by in vitro selection (Ducongé, F., and Toulmé, J. J. (1999) RNA5, 1605–1614). The high affinity sequences selected at physiological magnesium concentration (3 mm) were shown to form a loop-loop complex with the targeted TAR RNA. The stability of this complex depends on the aptamer loop closing "GA pair" as characterized by preliminary electrophoretic mobility shift assays. Thermal denaturation monitored by UV-absorption spectroscopy and binding kinetics determined by surface plasmon resonance show that the GA pair is crucial for the formation of the TAR-RNA aptamer complex. Both thermal denaturation and surface plasmon resonance experiments show that any other "pairs" leads to complexes whose stability decreases in the order AG > GG > GU > AA > GC > UA >> CA, CU. The binding kinetics indicate that stability is controlled by the off-rate rather than by the on-rate. Comparison with the complex formed with the TAR* hairpin, a rationally designed TAR RNA ligand (Chang, K. Y., and Tinoco, I. (1994) Proc. Natl. Acad. Sci. U. S. A. 91, 8705–8709), demonstrates that the GA pair is a key determinant which accounts for the 50-fold increased stability of the TAR-aptamer complex (K d= 2.0 nm) over the TAR-TAR* one (K d = 92.5 nm) at physiological concentration of magnesium. Replacement of the wild-type GC pair next to the loop of RNA I′ by a GA pair stabilizes the RNA I′-RNA II′ loop-loop complex derived from the one involved in the control of the ColE1 plasmid replication. Thus, the GA pair might be the preferred one for stable loop-loop interactions. human immunodeficiency virus type 1 nucleotide(s) surface plasmon resonance electrophoretic mobility shift assay resonance unit(s) trans-activation-responsive Intermolecular interactions between structured RNA play key roles in the regulation of gene expression. In Escherichia coli, the replication of the ColE1 plasmid is regulated by the interaction of two RNA transcripts, RNA I and RNA II, which fold as hairpins (1.Tomizawa J.I. Cell. 1986; 47: 89-97Abstract Full Text PDF PubMed Scopus (103) Google Scholar, 2.Eguchi Y. Tomizawa J.I. Cell. 1990; 60: 199-209Abstract Full Text PDF PubMed Scopus (110) Google Scholar). The interaction starts with base pairing between the complementary loops of these RNAs and leads to the formation of a double-stranded RNA along the entire length of RNA I, thus disrupting the RNA II hybridization with the template DNA required for replication (3.Tomizawa J.I. J. Mol. Biol. 1990; 212: 683-694Crossref PubMed Scopus (84) Google Scholar, 4.Masukata H. Tomizawa J.I. Cell. 1986; 44: 125-136Abstract Full Text PDF PubMed Scopus (120) Google Scholar). Even if the stability of these so-called "kissing" complexes is primarily based on loop complementarity (2.Eguchi Y. Tomizawa J.I. Cell. 1990; 60: 199-209Abstract Full Text PDF PubMed Scopus (110) Google Scholar), factors such as the orientation of the loops (5.Eguchi Y. Tomizawa J.I. J. Mol. Biol. 1991; 220: 831-842Crossref PubMed Scopus (90) Google Scholar), the loop closing base pair, and the sequence of each stem next to the loop (6.Gregorian Jr., R.S. Crothers D.M. J. Mol. Biol. 1995; 248: 968-984Crossref PubMed Scopus (72) Google Scholar), are crucial for stability. For instance, the loop inversion 5′ to 3′ induces a 350-fold increased stability of the RNA I-RNA II complex. In HIV-1,1 the dimerization of the genomic RNA involves the formation of a loop-loop complex between two structured regions (7.Paillart J.C. Westhof E. Ehresmann C. Ehresmann B. Marquet R. J. Mol. Biol. 1997; 270: 36-49Crossref PubMed Scopus (120) Google Scholar). The dimerization initiation site of HIV-1 RNA folds as a hairpin, which is closed by a noncanonical AA pair. The 9-nt-long loop contains a 6-nt self-complementary sequence flanked by two 5′ and one 3′ purines, which, together with loop complementarity, are crucial for the dimerization process (8.Jossinet F. Paillart J.C. Westhof E. Hermann T. Skripkin E. Lodmell J.S. Ehresmann C. Ehresmann B. Marquet R. RNA. 1999; 5: 1222-1234Crossref PubMed Scopus (89) Google Scholar). Non-Watson-Crick interactions in RNA molecules have been also reported in the viral RNA element bound by the Rev protein of HIV-1 (9.Bartel D.P. Zapp M.L. Green M.R. Szostak J.W. Cell. 1991; 67: 529-536Abstract Full Text PDF PubMed Scopus (348) Google Scholar), in GRNA tetraloops (10.Lehnert V. Jaeger L. Michel F. Westhof E. Chem. Biol. 1996; 3: 993-1009Abstract Full Text PDF PubMed Scopus (276) Google Scholar, 11.Jestin J.L. Deme E. Jacquier A. EMBO J. 1997; 16: 2945-2954Crossref PubMed Scopus (37) Google Scholar), tRNAs (12.Leontis, N. B. (2000) Q. Rev. Biophysics, in pressGoogle Scholar, 13.Dirheimer G. Keith G. Dumas P. Westhof E. Söll D. RajBhandary U. tRNA: Structure, Biosynthesis, and Function. American Society for Microbiology, Washington, D.C.1995Google Scholar, 14.Tinoco I. Gesteland R.F. Atkins J.F. The RNA World. Cold Spring Harbor Laboratory Press, Cold Spring, Harbor, NY1993Google Scholar), and tandem mismatches within duplexes (15.SantaLucia Jr., J. Turner D.H. Biochemistry. 1993; 32: 12612-12623Crossref PubMed Scopus (175) Google Scholar, 16.Heus H.A. Wijmenga S.S. Hoppe H. Hilbers C.W. J. Mol. Biol. 1997; 271: 147-158Crossref PubMed Scopus (53) Google Scholar, 17.Biswas R. Sundaralingam M. J. Mol. Biol. 1997; 270: 511-519Crossref PubMed Scopus (44) Google Scholar). All these results indicate that interactions other than canonical base pairs contribute to the structural diversity displayed by RNAs, which, as a matter of fact, is crucial for activity. The contribution of such noncanonical interactions can actually be conveniently explored by in vitro selection, since neither the structure of the target nor the interactions between the target and the interacting aptamer need to be known (18.Ellington A.D. Conrad R. Bio/Technol. Annu. Rev. 1995; 1: 185-215Crossref PubMed Scopus (53) Google Scholar, 19.Gold L. Polisky B. Uhlenbeck O. Yarus M. Annu. Rev. Biochem. 1995; 64: 763-797Crossref PubMed Scopus (747) Google Scholar). This strategy was successfully used in our laboratory to identify DNA aptamers against DNA (20.Mishra R.K. Toulmé J.J. C. R. Acad. Sci. Paris. 1994; 317: 977-982PubMed Google Scholar, 21.Mishra R.K. Le Tinévez R. Toulmé J.J. Proc. Natl. Acad. Sci. U. S. A. 1996; 93: 10679-10684Crossref PubMed Scopus (23) Google Scholar, 22.Boiziau C. Dausse E. Mishra R. Ducongé F. Toulmé J.J. Antisense Nucleic Acid Drug D. 1997; 7: 369-380Crossref PubMed Scopus (12) Google Scholar) and RNA hairpin structures (23.Boiziau C. Dausse E. Yurchenko L. Toulmé J.J. J. Biol. Chem. 1999; 274: 12730-12737Abstract Full Text Full Text PDF PubMed Scopus (104) Google Scholar). Recently, RNA aptamers specific for the trans-activation-responsive (TAR) RNA (24.Karn J. J. Mol. Biol. 1999; 293: 235-254Crossref PubMed Scopus (391) Google Scholar) were selected by in vitro selection (25.Ducongé F. Toulmé J.J. RNA. 1999; 5: 1605-1614Crossref PubMed Scopus (118) Google Scholar). The TAR RNA element is a 59-nt-long imperfect stem loop structure located at the 5′ end of the retroviral RNA. A 3-nt bulge in the upper part of the hairpin constitutes part of the binding site of the viral protein Tat, which recruits cyclin T1. Together with additional TAR-bound cellular proteins, this complex prevents abortion of the transcription of the retroviral genome. Therefore, TAR plays a key role in the life cycle of HIV-1 and constitutes a valid target for the development of ligands, which could inhibit its interaction with viral and cellular proteins, thus ultimately preventing the development of the virus. The isolated high affinity anti-TAR aptamers were shown to fold as imperfect hairpins and form kissing complexes with the targeted RNA at physiological magnesium concentration. The apical loop of all these aptamers corresponds to the 5′-GUCCCAGA-3′ consensus sequence, the six central bases of which are complementary to the entire TAR loop whereas a conserved "GA pair" closes the loop. As forColE1 RNA I and RNA II, and HIV-1 dimerization initiation site RNA motif, interactions other than loop complementarity are crucial for stability. Indeed, several mutations of the GA pair that closes the loop of the identified RNA aptamers decrease the stability of the TAR-aptamer complex, as shown by preliminary electrophoretic mobility shift assays (EMSA). In the work presented herein, the role of this GA pair was investigated at physiological concentration of magnesium by systematically mutating the loop closing pair of the aptamer. The effects of these mutations on the stability of the TAR RNA-aptamer complex were analyzed by thermal denaturation monitored by UV-absorption spectroscopy (26.Breslauer K.J. Agrawal S. Protocols for Oligonucleotide Conjugates. 26. Humana Press Inc., 1994: 347-372Google Scholar, 27.Puglisi J.D. Tinoco Jr., I. Methods Enzymol. 1989; 180: 304-325Crossref PubMed Scopus (658) Google Scholar). The binding kinetics were determined by using surface plasmon resonance (SPR). This physical phenomenon is used to follow in real time the interaction between a molecule in a continuous flow and an immobilized one (28.Jonsson U. Fagerstam L. Ivarsson B. Johnsson B. Karlsson R. Lundh K. Lofas S. Persson B. Roos H. Ronnberg I. et al.BioTechniques. 1991; 11: 620-627PubMed Google Scholar). Numerous studies have been published on protein-protein interactions, protein-ligand interactions, and protein-nucleic acid interactions, but a much reduced number of investigations of nucleic acid-nucleic acid interactions are available (29.Crouch R.J. Wakasa M. Haruki M. Methods Mol. Biol. 1999; 118: 143-160PubMed Google Scholar, 30.Morton T.A. Myszka D.G. Methods Enzymol. 1998; 295: 268-294Crossref PubMed Scopus (267) Google Scholar, 31.Schuck P. Curr. Opin. Biotechnol. 1997; 8: 498-502Crossref PubMed Scopus (142) Google Scholar). As a matter of fact, no RNA-RNA complexes have been analyzed up to now. Our results demonstrate that the aptamer loop closing GA pair is crucial for the stability of the TAR-RNA aptamer complex once formed. This likely explains the higher stability of the loop-loop complex formed by TAR with the aptamer, at physiological magnesium concentration, compared with the one formed with the rationally designed hairpin, TAR*, whose loop is closed by a UA pair (32.Chang K.Y. Tinoco I. Proc. Natl. Acad. Sci. U. S. A. 1994; 91: 8705-8709Crossref PubMed Scopus (93) Google Scholar). The increased stability of a loop-loop complex formed between RNA II′ and a RNA I′ mutant in which the GC pair next to loop was replaced by GA suggests that closing GA pair could be preferred for kissing complexes. All RNA molecules including the biotinylated TAR RNA were synthesized on an Expedite 8908 synthesizer and purified by electrophoresis on denaturing polyacrylamide gels. The pure samples were desalted on Sephadex G-25 spin columns. To avoid repeated thawing and freezing of the stock solutions, the samples were aliquoted at a volume and a concentration suitable for each experiment and stored at −20 °C. Before the experiments, each RNA sample used was heated at 95 °C during 1 min and then put on ice for 10 min to avoid the formation of intermolecular complexes. Thermal denaturation of RNA complexes in 20 mm sodium cacodylate buffer, pH 7.3, at 20 °C, with 140 mm potassium chloride, 20 mm sodium chloride and 3 mm magnesium chloride (R′ buffer) was monitored on a Cary 1 spectrophotometer interfaced with a Peltier effect device that controls temperature within ±0.1 °C. Denaturation of the samples was achieved by increasing the temperature at 0.4 °C/min from 5 °C to 90 °C and was followed at 260 nm. A cuvette that contained R′ buffer was used as the reference. Except cacodylate, which replaced the temperature-sensitive HEPES, the buffer used for these thermal denaturation experiments was equivalent to the one used during the in vitro selection process (25.Ducongé F. Toulmé J.J. RNA. 1999; 5: 1605-1614Crossref PubMed Scopus (118) Google Scholar). As the absorbance of the TAR RNA at 260 nm is too large to allow accurate monitoring of the absorption change resulting from the denaturation of the bimolecular complex between the HIV-1 RNA and the RNA aptamer, the experiments were carried on with miniTAR, a 27-mer oligonucleotide, instead of the entire TAR hairpin, which is 59 nt. RNA samples were prepared at 1 μm final concentration in the mixture. They were mixed at room temperature and allowed to interact 30 min before cooling down to 5 °C. The experiment then started after 1 h at this temperature. The enthalpy change, ΔH, for the formation of the bimolecular complex, was deduced from the total RNA concentration dependence of the Tm according to Equation1. 1Tm=RΔHln[ARN] total+ΔS−Rln4ΔHEquation 1 The number of magnesium ions that bind upon formation of the complex, ΔMg2+, was determined from the ion concentration dependence of the melting temperature, Tm , according to Equation 2. δ(1/Tm)δln[Mg2+]=ΔMg2+RΔHEquation 2 SPR experiments were performed on a BIAcore 2000 apparatus (Biacore AB, Sweden) running with the BIAcore 2.1 software. Biotinylated TAR RNA, 59 nt long, was immobilized on CM5 sensorchips coated with streptavidin according to the procedure described in the BIA applications handbook. In these conditions, 5000 resonance units (RU) of streptavidin (Sigma), equivalent to 5 ng/mm2, were immobilized on the chip which subsequently was allowed to equilibrate at 23 °C, the temperature of the in vitro selection, in the selection buffer: 20 mm HEPES, pH 7.3, at 20 °C containing 20 mmsodium acetate, 140 mm potassium acetate, and 3 mm magnesium acetate (R buffer). Biotinylated TAR RNA (10–50 nm) was prepared in this buffer and then injected at a flow rate of 5 μl/min. The injection was stopped as soon as 500–600 RU of bound TAR RNA was reached. This amount was shown to be appropriate to keep the pseudo-first order kinetic condition and to allow good reliability of recorded sensorgrams (RU versustime) in terms of signal to noise ratio. One noncoated or one streptavidin-coated flow-cell was used to check for nonspecific binding of RNA aptamers. The signals from these control channels served as base lines and were subtracted to the RU change observed when an injected RNA aptamer interacts with the immobilized TAR RNA. SL1 RNA, a nonrelevant hairpin from the hepatitis C virus RNA, was used as a negative control (33.Blight K.J. Rice C.M. J. Virol. 1997; 71: 7345-7352Crossref PubMed Google Scholar). The sensorchip surface was successfully regenerated with three 5-μl pulses of 25% formamide, followed by one 5-μl pulse of distilled water and finally one 10-μl pulse of R buffer. Nonlinear regression analysis of single sensorgrams at five concentrations, at least, of injected RNAs was used to determine the kinetic parameters of the complex formation. The data were analyzed with the BIA evaluation 2.2.4 software, assuming a pseudo-first order model, according to Equations Equation 3, Equation 4, Equation 5, for the association and dissociation phases, respectively, where R is the signal response, R max the maximum response level,C the molar concentration of the injected RNA molecule,k on the association rate constant, andk off the dissociation rate constant. dRdt=konC(Rmax−R)−koffREquation 3 dRdt=−koffREquation 4 To check for self-consistency of data,k obs, derived from nonlinear analysis, was plotted as a function of the RNA concentration according to Equation5. kobs=konC+koffEquation 5 The RNA molecules used for this study are shown in Fig.1. MiniTAR (27 nt long), the upper half part of TAR, was shown to be the mininal domain necessary and sufficient for responsiveness in vivo (34.Jakobovits A. Smith D.H. Jakobovits E.B. Capon D.J. Mol. Cell. Biol. 1988; 8: 2555-2561Crossref PubMed Google Scholar). It interacts with the anti-TAR selected aptamers without changes in the affinity compared with the full-length TAR (35.Ducongé F. Ph.D thesis. Université Victor Segalen, Bordeaux, France1999Google Scholar). R-0624(GA) is the aptamer of highest affinity identified by in vitro selection against TAR, with the consensus motif 5′-GUCCCAGA-3′, LR-068, in the apical loop. The six central bases of the consensus sequence are complementary to the entire TAR loop. TAR* is a hairpin rationally designed to interact with TAR (32.Chang K.Y. Tinoco I. Proc. Natl. Acad. Sci. U. S. A. 1994; 91: 8705-8709Crossref PubMed Scopus (93) Google Scholar). Its loop is fully complementary to the TAR one. RNA I′ and RNA II′ are two structured RNAs derived from the two RNA transcripts, RNA I and RNA II, involved in the control of the ColE1 plasmid regulation. The sequences of RNA I′ and RNA II′ were modified in the stem to avoid the formation of an extended duplex, as seen with the biological RNAs once the kissing complex is formed (6.Gregorian Jr., R.S. Crothers D.M. J. Mol. Biol. 1995; 248: 968-984Crossref PubMed Scopus (72) Google Scholar). The derivative of UV melting curves of miniTAR with R-0624(GA) aptamer, with TAR* RNA and with LR-068 are reported in Fig.2. The melting profiles obtained with mixtures of miniTAR and R-0624(GA) RNA at two different concentrations in R′ buffer display two transitions (Fig.2 A). Only one transition is observed for the melting profiles of the RNA hairpins alone. On diluting the miniTAR and R-0624(GA) aptamer mixture 4-fold (from 2 μmto 0.5 μm), the Tm (the maximum of the derivative plots) of the lower transition decreases from 47.5 °C to 40 °C as expected for a bimolecular complex, whereas theTm of the higher transition remains unchanged and thus can be ascribed to the melting of the RNAs alone. We then compared the stability of the miniTAR and R-0624(GA) aptamer mixture to that of two reference complexes: miniTAR with either TAR* RNA or LR-068 (Fig.2 B). Under the ionic conditions used for the in vitro selection, Tm for the complex with TAR* is equal to 30.7 °C (Table I),i.e. 16.8 °C below that of the miniTAR-R-0624(GA) complex. Finally,Tm for the complex with LR-068, the 8-mer RNA 5′-GUCCCAGA-3′ consensus motif of the aptamers, is equal to 20.3 °C. Clearly, interaction of miniTAR with the aptamer gives rise to the most stable bimolecular complex. On one hand, the higher stability of the TAR-aptamer complex over that of the 8-mer RNA can be ascribed to the hairpin structure of the ligand as previously demonstrated (36.Grosjean H. Soll D.G. Crothers D. J. Mol. Biol. 1976; 103: 499-519Crossref PubMed Scopus (172) Google Scholar). On the other hand, the comparison between TAR* and the selected aptamer strongly suggests that the sequence outside the loop plays a crucial role in the extra stability displayed by TAR-aptamer complexes compared with the TAR-TAR* one.Table IMelting temperature, Tm, of RNA complexes and RNAs aloneRNAT mMiniTAR-RNA complexRNA alone°C°CR-0624(GA)47.3 ± 0.370.8 ± 0.3R-0624(AG)42.8 ± 0.470.7 ± 0.7R-0624(GG)37.0 ± 0.072.3 ± 0.0R-0624(GU)32.9 ± 0.171.4 ± 0.0R-0624(AA)31.5 ± 0.665.1 ± 0.6R-0624(GC)31.4 ± 0.681.5 ± 0.0R-0624(UA)29.9 ± 0.675.3 ± 0.5R-0624(CA)21.0 ± 0.070.3 ± 0.4R-0624(CU)16.8 ± 1.173.0 ± 0.1Tar*30.7 ± 0.666.1 ± 0.8R-0616(UA)31.4 ± 0.562.9 ± 0.4LR-06820.3 ± 0.4ND aND, not determined.MiniTAR69.9 ± 0.2RNA I′-RNA II′ complexRNA I′ (GC)ND75.8 ± 0.0RNA I′ (GA)23.8 ± 0.870.0 ± 0.0RNA II′70.4 ± 0.6The experiments with miniTAR either with TAR*, the RNA aptamers or the truncated versions were performed in R′ buffer at 1 μmeach RNA, those with RNA I′ and RNAII′ in R′ buffer + 7 mm Mg2+ at 2 μm each RNA.T m is the average and standard deviation of two or three experiments.a ND, not determined. Open table in a new tab The experiments with miniTAR either with TAR*, the RNA aptamers or the truncated versions were performed in R′ buffer at 1 μmeach RNA, those with RNA I′ and RNAII′ in R′ buffer + 7 mm Mg2+ at 2 μm each RNA.T m is the average and standard deviation of two or three experiments. The stability of RNA structures and complexes is well known to depend on magnesium ion. We then wanted to address the question whether the differential behavior between TAR* and the aptamer would originate in the number of associated Mg2+ ions. This can be achieved by measuring the variation of Tm as a function of the Mg2+ concentration if the enthalpy change, ΔH, for the transition is known. The enthalpy change for the formation of the complex, ΔH, was deduced from the slope of the total RNA concentration dependence of theTm (Fig. 3), according to Equation 1. In these experiments, the concentration of Mg2+ was decreased from 3 mm in R′ buffer to 1 and 0.1 mm, for the miniTAR-TAR* and miniTAR-aptamer complexes, respectively, to accurately measure theTm values of bimolecular complexes in the total RNA concentration range chosen, 0.5–16 μm, with no interference with the melting of the hairpins alone. Under these conditions, ΔH is equal to −42.8 ± 1.4 kcal/mol and −39.2 ± 1.3 kcal/mol for miniTAR-R-0624(GA) and miniTAR-TAR* complexes, respectively. The variation ofTm as a function of the Mg2+concentration is presented in Fig. 4. For both complexes, linearity of 1/Tmversus ln[Mg2+] plots indicates a site binding mode of the magnesium ion. The number of ions, ΔMg2+, which bind was deduced according to Equation 2 using the enthalpy change. This gives ΔMg2+ = 1.7 ± 0.1 and 1.4 ± 0.1 for the complexes formed with R-0624(GA) and TAR*, respectively. It demonstrates that the different stability of these complexes does not originate in a difference in the number of magnesium ions that each complex binds. From this ΔH and theTm values measured at 3 mm magnesium (Table I), the equilibrium dissociation constant,K d, in solution at 23 °C, for both complexes can be easily determined. K d, under the in vitro selection conditions, is equal to 2.0 ± 0.4 nm and 92.5 ± 5.2 nm for miniTAR-R-0624(GA) and miniTAR-TAR* complexes, respectively. Clearly, under this condition, the miniTAR-aptamer complex is more stable than the one formed with TAR*, the rationally designed RNA.Figure 4Dependence of T m on the concentration of Mg2+ for miniTAR complexes. The experiments were performed using 1 μm amount of each RNA. Complex with TAR* (○) or with R-0624(GA) (▪) is shown. Linear fits were calculated according to Equation 2.View Large Image Figure ViewerDownload Hi-res image Download (PPT) The base pair closing the loop was a marked difference between TAR* and the aptamer molecule. Then, in a second set of experiments, the closing GA pair was mutated to evaluate its role on the stability of the miniTAR-aptamer complex. The derivatives of the UV melting curves for complexes with three aptamer variants, namely R-0624(GA), R-0624(CU), and R-0624(AA), are presented in Fig. 5. Similar measurements have been carried out for other combinations, and Tm values for all complexes and variants alone are listed in Table I. Clearly, any change of the closing pair induces a decrease inTm that reflects a decrease in the stability of the miniTAR-aptamer complex. SPR was used to follow, in real time, the interaction of the immobilized full-length TAR RNA on streptavidin-coated sensorchip with various RNA hairpins. Sensorgrams, obtained when R-0624(GA) or TAR* analytes were injected over the sensorchip surface at two different concentrations of magnesium ion, are reported in Fig.6. In all cases, as expected for a pseudo-first order model, the dissociation phase does not show significant dependence on the concentration of the injected analyte whereas the association phase increases with it. Furthermore, these kinetics fulfill pseudo-first order conditions as checked from linearity of plots of the observed rate constant,k obs, versus the analyte concentration (insets). R-0624(GA) aptamer binds to TAR RNA at either 3 or 10 mm magnesium, as seen in Fig.6 (A and B, respectively). In contrast, whereas TAR* binds also at 10 mm (Fig. 6 D), it shows a poor affinity at 3 mm (Fig. 6 C). The rate constants, k on and k off, deduced from these sensorgrams are listed in TableII. For R-0624(GA), the on-rate changes from 6.3 × 104m−1 s−1to 17 × 104m−1s−1 upon increasing the concentration of magnesium from 3 to 10 mm. The off-rate remains constant and is equal to about 10−3s−1. For TAR*, at 3 mm magnesium, the rate constants cannot be reasonably determined. At 10 mm, k on andk off are equal to 38 × 104m−1 s−1and 5.3 × 10−3s−1, respectively. Also listed in Table II are the equilibrium dissociation constants deduced from these kinetic measurements, K d, equal tok off/k on, which confirms that the TAR-aptamer complex is much more stable in the in vitro selection buffer than the one with the rationally designed TAR* molecule, in agreement with EMSA (25.Ducongé F. Toulmé J.J. RNA. 1999; 5: 1605-1614Crossref PubMed Scopus (118) Google Scholar). As expected, at 10 mm magnesium, i.e. the concentration used to investigate the TAR-TAR* complex (32.Chang K.Y. Tinoco I. Proc. Natl. Acad. Sci. U. S. A. 1994; 91: 8705-8709Crossref PubMed Scopus (93) Google Scholar), the stability of both complexes increases. The stabilizing effect observed on the complex with R-0624(GA) resides essentially in the association step, as expected for larger screening of electrostatic repulsions.Table IIEquilibrium and rate constants for TAR binding to R-0624 or TAR*BufferRNAk onk offK d(BIAcore)K d(EMSA)× 10 4 m −1 s −1× 10 −3 s −1nmnmRR-06246.3 ± 0.61.1 ± 0.117 ± 320TAR*ND aND, not determined.NDND>1000R + 7 mm Mg2+R-062417 ± 10.93 ± 0.15.4 ± 0.86TAR*38 ± 75.3 ± 0.114.5 ± 310The experiments were performed either in R buffer or in R buffer + 7 mm Mg2+. K d(BIAcore)was calculated as k off/k on andK d(EMSA) was derived from electrophoretic mobility shift assays (Ducongé & Toulmé, 1999).k on, k off, andK d are the averages and standard deviations of at least five sensorgrams where R-0624 or TAR* was injected at five different concentrations at 20 μl/min at 23 °C.a ND, not determined. Open table in a new tab The experiments were performed either in R buffer or in R buffer + 7 mm Mg2+. K d(BIAcore)was calculated as k off/k on andK d(EMSA) was derived from electrophoretic mobility shift assays (Ducongé & Toulmé, 1999).k on, k off, andK d are the averages and standard deviations of at least five sensorgrams where R-0624 or TAR* was injected at five different concentrations at 20 μl/min at 23 °C. The role of the GA closing pair of R-0624 RNA was further examined by SPR in R buffer (3 mm magnesium). Mutants of the closing pair were injected, and RU variations that resulted from complex formation were monitored as a function of time. The rate constants, k on and k off, deduced from nonlinear regression analysis of sensorgrams are listed in Table III as well as the equilibrium dissociation constant, K d. Except for the AG inversion, which is equivalent to the wild-type aptamer in terms of stability, with a K d of about 20 nm, all other mutations have a negative effect on the stability of the TAR-RNA complex. CU and CA mutations destabilize the TAR-aptamer complex to such a degree that the rate constants cannot be determined. The effects of the other mutations are in between. Closer analysis shows that the equilibrium constant is mainly controlled by the off-rate, which increases when the complex is destabilized, whereas the on-rate shows limited variation.Table IIIEffects of mutations of the loop closing GA pair of the aptamer on equilibrium and rate constants for TAR bindingRNAk onk offK d(BIAcore)K d(EMSA)× 10 4 m −1 s −1× 10 −3 s −1nmnmR-0624(GA)6.3 ± 0.61.1 ± 0.117 ± 318 ± 2R-0624(AG)6.8 ± 0.91.5 ± 0.122 ± 232 ± 9R-0624(GG)7.9 ± 0.44.1 ± 0.152 ± 263 ± 15R-0624(GU)8.5 ± 1.49.1 ± 0.2107 ± 14133 ± 20R-0624(AA)5.7 ± 0.56.4 ± 0.1122 ± 12340 ± 80R-0624(GC)10.5 ± 0.318.5 ± 0.8167 ± 22264 ± 130R-0624(UA)6.3 ± 0.313.0 ± 0.1206 ± 1440 ± 130R-0624(CA)ND aND, not