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• W2023363067 abstract "To investigate the relationship between RNA folding and ribozyme catalysis, we have carried out a detailed kinetic analysis of four structural derivatives of the hairpin ribozyme. Optimal and suboptimal (wild-type) substrate sequences were studied in conjunction with stabilization of helix 4, which supports formation of the catalytic core. Pre-steady-state and steady-state kinetic studies strongly support a model in which each of the ribozyme variants partitions between two major conformations leading to active and inactive ribozyme· substrate complexes. Reaction rates for cleavage, ligation, and substrate binding to both ribozyme conformations were determined. Ligation rates (3 min−1) were typically 15-fold greater than cleavage rates (0.2 min−1), demonstrating that the hairpin ribozyme is an efficient RNA ligase. On the other hand, substrate binding is very rapid (kon = 4 × 108m−1 min−1), and the ribozyme· substrate complex is very stable (KD< 25 pm; koff < 0.01 min−1). Stabilization of helix 4 increases the proportion of RNA molecules folded into the active conformation, and enhances substrate association and ligation rates. These effects can be explained by stabilization of the catalytic core of the ribozyme. Rigorous consideration of conformational isomers and their intrinsic kinetic properties was necessary for development of a kinetic scheme for the ribozyme-catalyzed reaction. To investigate the relationship between RNA folding and ribozyme catalysis, we have carried out a detailed kinetic analysis of four structural derivatives of the hairpin ribozyme. Optimal and suboptimal (wild-type) substrate sequences were studied in conjunction with stabilization of helix 4, which supports formation of the catalytic core. Pre-steady-state and steady-state kinetic studies strongly support a model in which each of the ribozyme variants partitions between two major conformations leading to active and inactive ribozyme· substrate complexes. Reaction rates for cleavage, ligation, and substrate binding to both ribozyme conformations were determined. Ligation rates (3 min−1) were typically 15-fold greater than cleavage rates (0.2 min−1), demonstrating that the hairpin ribozyme is an efficient RNA ligase. On the other hand, substrate binding is very rapid (kon = 4 × 108m−1 min−1), and the ribozyme· substrate complex is very stable (KD< 25 pm; koff < 0.01 min−1). Stabilization of helix 4 increases the proportion of RNA molecules folded into the active conformation, and enhances substrate association and ligation rates. These effects can be explained by stabilization of the catalytic core of the ribozyme. Rigorous consideration of conformational isomers and their intrinsic kinetic properties was necessary for development of a kinetic scheme for the ribozyme-catalyzed reaction. Since the first description of a catalytically active RNA molecule (1Cech T.R. Zaug A.J. Grabowski P.J. Cell. 1981; 27: 487-496Google Scholar), much effort has been focused toward elucidating the molecular mechanisms of ribozyme catalysis. Valuable information has emerged from detailed kinetic and thermodynamic analyses of intramolecular and intermolecular reactions catalyzed by several naturally occurring ribozymes, including self-splicing group I introns (2Herschlag D. Cech T.R. Biochemistry. 1990; 29: 10159-10171Google Scholar, 3Bevilacqua P.C. Kierkez R. Johnson K.A. Turner D.H. Science. 1992; 258: 1355-1358Google Scholar, 4Bevilacqua P.C. Sugimoto N. Turner D.H. Biochemistry. 1996; 35: 648-658Google Scholar, 5Golden B.L. Cech T.R. Biochemistry. 1996; 35: 3754-3763Google Scholar, 6Mei R. Herschlag D. Biochemistry. 1996; 35: 5796-5809Google Scholar) and group II introns (7Pyle A.M. Green J.B. Biochemistry. 1994; 33: 2716-2725Google Scholar, 8Michels Jr., W.J. Pyle A.M. Biochemistry. 1995; 34: 2965-2977Google Scholar, 9Daniels D.L. Michels Jr., W.J. Pyle A.M. J. Mol. Biol. 1996; 256: 31-49Google Scholar), ribonuclease P (10Beebe J.A. Fierke C.A. Biochemistry. 1994; 33: 10294-10304Google Scholar, 11Beebe J.A. Kurz J.C. Fierke C.A. Biochemistry. 1996; 35: 10493-10505Google Scholar), hammerhead ribozymes (12Hertel K.J. Herschlag D. Uhlenbeck O.C. Biochemistry. 1994; 33: 3374-3385Google Scholar, 13Hertel K.J. Uhlenbeck O.C. Biochemistry. 1995; 34: 1744-1749Google Scholar). and hairpin ribozymes (14Hegg L.A. Fedor M.J. Biochemistry. 1995; 34: 15813-15828Google Scholar). It is widely accepted that the folded structure of RNA is critical for its catalytic activity. However, few studies have addressed the problem of how differences in ribozyme folding may affect individual steps of the reaction pathway. One major complication in kinetic analysis of ribozymes results from the ability of most RNA molecules to fold into multiple conformations (15Uhlenbeck O.C. RNA. 1995; 1: 4-6Google Scholar). We believe that the study of conformationally heterogeneous ribozymes is important because it represents a direct and realistic approach to the problem of RNA structure and function. As a model to investigate the relationship between RNA structure and kinetic behavior, we are studying the hairpin ribozyme. This ribozyme is a relatively small RNA molecule (50 nucleotides, 17 kDa) derived from the minus strand of the satellite RNA associated with tobacco ringspot virus (16Feldstein P.A. Buzayan J.M. Bruening G. Gene ( Amst .). 1989; 82: 53-61Google Scholar, 17Hampel A. Tritz R. Biochemistry. 1989; 28: 4929-4933Google Scholar, 18Haseloff J. Gerlach W.L. Gene. 1989; 82: 43-52Google Scholar). It acts as a reversible, site-specific endoribonuclease, cleaving RNA substrates at a NpG linkage using a transesterification mechanism to generate products containing 5′-hydroxyl and 2′,3′-cyclic phosphate termini or ligating molecules with such end structures. The secondary structure of the ribozyme· substrate complex, as well as the nucleotides and functional groups required for catalysis, has been elucidated through mutational studies, phylogenetic analysis, and in vitroselection experiments (for review, see Ref. 19Burke J.M. Nucleic Acids & Mol. Biol. 1994; 8: 105-118Google Scholar). The substrate interacts with the ribozyme through two intermolecular helices, H1 and H2 (see Fig. 1 A), separated by a symmetrical loop (loop A) composed of four nucleotides in both substrate and ribozyme. Within the ribozyme, two intramolecular helices (H3 and H4) are separated by a large asymmetrical loop (loop B). Nucleobases essential for catalysis are concentrated within loops A and B. Chemical modification and linker-insertion experiments have led to the hypothesis that loop A and loop B establish tertiary interactions for the ribozyme catalytic core to be formed, therefore implying a sharp bend between helix 2 and helix 3 (for review, see Ref. 20Burke J.M. Butcher S.E. Sargueil B. Nucleic Acids & Mol. Biol. 1996; 10: 129-143Google Scholar). This hypothesis has received further support from experiments which demonstrated that activity can be reconstituted following separation of the A and B domains (21Butcher S.E. Heckman J.E. Burke J.M. J. Biol. Chem. 1995; 270: 29648-29651Google Scholar). Loop B and its flanking helices constitute an independent folding domain, as indicated by cross-linking and chemical modification studies (22Butcher S.E. Burke J.M. J. Mol. Biol. 1994; 244: 52-63Google Scholar, 23Butcher S.E. Burke J.M. Biochemistry. 1994; 33: 992-999Google Scholar). Although, to a first approximation, the sequence of the helices is not relevant for ribozyme activity (24Berzal-Herranz A. Joseph S. Chowrira B.M. Butcher S.E. Burke J.M. EMBO J. 1993; 12: 2567-2574Google Scholar), we found that extension of helix 4 produces a significant increase in catalytic proficiency, probably through stabilizing the active tertiary structure of loop B as measured by photocross-linking (25Sargueil B. Pecchia D.B. Burke J.M. Biochemistry. 1995; 34: 7739-7748Google Scholar). To achieve a better understanding of structure-function relationships in the hairpin ribozyme system, we have carried out a detailed kinetic analysis of different structural variants of this ribozyme. First, the effect of extending helix 4 was analyzed since its length is likely to affect the stability of the flanking catalytic core. Second, hairpin ribozymes with different substrate specificity were also examined because the naturally occurring substrate is conformationally heterogeneous (26Chowrira B.M. Burke J.M. Biochemistry. 1991; 30: 8518-8522Google Scholar). 1Heckman et al., unpublished observations. 1Heckman et al., unpublished observations. Therefore, four derivatives of the hairpin ribozyme that combine these two modifications, were studied. As shown in Fig. 1, the original helix 4 was extended with three extra base pairs and a stable GUAA tetraloop, and the sequence of the substrate and ribozyme were varied to avoid substrate self-complementarity, as described in Butcher et al. (21Butcher S.E. Heckman J.E. Burke J.M. J. Biol. Chem. 1995; 270: 29648-29651Google Scholar). The four resulting hairpin ribozymes were assayed in combination with their cognate substrates. Substrate specificity will be indicated as original sequence (wt) 2The abbreviations used are: wt, wild-type or original sequence; SV5, modified sequence; EH4, extended helix 4. 2The abbreviations used are: wt, wild-type or original sequence; SV5, modified sequence; EH4, extended helix 4. or modified sequence (SV5), and the presence of an extended helix 4 will be referred to as EH4. This comparative analysis has provided new data on the relationship between RNA folding and catalysis by detecting two inherent conformational states of the hairpin ribozyme and interpreting the resulting biphasic kinetics in terms of individual rate constants. These results provide important insights into folding of the hairpin ribozyme and illustrate how structural diversity can be reflected in kinetic behavior. We expect that our results will also be useful for the rational design of new ribozymes with more homogeneous folding and improved catalytic efficiency. DNA templates for ribozyme transcription and ribozyme substrates (see Fig. 1) were synthesized on an Applied Biosystems 392 DNA/RNA synthesizer using standard DNA and RNA phosphoramidite chemistry. The sequence of the DNA oligonucleotide complementary to the wt RNA substrate was 5′-CCAAACAGGACTGTCGGTTG-3′. Ribozymes were synthesized by transcribing partially duplex synthetic DNA templates with T7 RNA polymerase, basically as described (27Milligan J.F. Uhlenbeck O.C. Methods Enzymol. 1989; 180: 51-62Google Scholar). All DNA and RNA molecules were purified by polyacrylamide gel electrophoresis as described (25Sargueil B. Pecchia D.B. Burke J.M. Biochemistry. 1995; 34: 7739-7748Google Scholar). In addition, RNA products of solid-phase synthesis were purified by reversed-phase high pressure liquid chromatography. RNA substrates were 5′-end-labeled with [γ-32P]ATP and T4 polynucleotide kinase. For RNA ligations in cis, ribozymes were internally labeled with [α-32P]CTP during transcription. All reactions were carried out in a reaction buffer containing 50 mm Tris-HCl, pH 7.5, and 12 mm MgCl2 at 25 °C. Ribozyme and substrate RNAs were preincubated separately for 10 min at 37 °C in reaction buffer. Complete denaturation of ribozymes was avoided to prevent formation of ribozyme dimers (23Butcher S.E. Burke J.M. Biochemistry. 1994; 33: 992-999Google Scholar). The solutions were then allowed to equilibrate for 10 min at 25 °C. Reactions were initiated by mixing equal volumes of solutions containing the ribozyme and substrate. Aliquots of the reaction (10 μl) were removed and quenched with an equal volume of loading buffer (15 mm EDTA, 97% formamide). Samples were analyzed in 20% (cleavage andtrans-ligation assays) or 6% (cis-ligation assays) polyacrylamide-urea (7 m) gel electrophoresis. Radioactive bands were quantified using a Bio-Rad GS-525 radioimaging system. Cleavage reactions were carried out with 200 nm ribozyme and less than 1 nm 5′-32P-substrate, unless otherwise indicated. No change either in the rate or in the extent of cleavage was observed at higher ribozyme concentrations, indicating that 200 nm ribozyme is saturating (data not shown). The fraction of substrate cleaved was plotted versus time and fitted to single- or double-exponential equations. The single-exponential equation was, Fraction reacted=−A·e−r·t+BEquation 1 where A and r stand for the amplitude and the rate of the exponential time course, respectively. The double-exponential equation was, Fraction reacted=−A1·e−r1·t−A2·e−r2·t+BEquation 2 where A1 and A2represent the amplitudes of the biphasic time course, andr1 and r2 stand for the corresponding rates. B represents the end point of the cleavage reaction and was typically between 0.6 and 0.9. These parameters were estimated by nonlinear regression analysis using the Marquardt-Levenberg algorithm (Sigma Plot 5.0 software and Origin/Microcal software). The standard error for the fitted parameters was typically less than 10%. Experimental error from independent measurements was less than 50% for wt ribozymes and less than 25% for SV5 ribozymes. Experiments carried out side by side were highly reproducible (less than 5% error). It is important to mention that experimental errors never altered the qualitative behavior of a reaction, which is, reactions that were best fitted with a double-exponential equation remained consistently biphasic despite the experimental error. The same was true for monophasic reactions. Cleavage reactions were carried out with 1–10 nm substrate and 0.1 nm ribozyme. Reactions were incubated at different times to obtain initial velocities for each substrate concentration. In some cases, higher substrate concentrations (1 μm) were used to evaluate the linearity of the product formation velocity at long reaction times (up to 5 h). Kinetic parameters were obtained by fitting the data to the Michaelis-Menten equation, kobs=kcatSS+KMEquation 3 where S stands for total substrate concentration. The steady-state parameters kcat andKm were estimated by nonlinear regression analysis as described above. For reactions in cis, ligation assays utilize self-cleaving molecules in which the 5′-end of the substrate is tethered through a short linker (five consecutive cytidines) to the 3′-end of the ribozyme (28Berzal-Herranz A. Joseph S Burke J.M. Genes Dev. 1992; 6: 129-134Google Scholar). These molecules were obtained by transcription from synthetic DNA templates in the presence of [α-32P]CTP. RNA self-cleavage takes place during transcription, and the larger product (ribozyme tethered to the 5′ cleavage product containing a 2′,3′-cyclic phosphate) was gel-purified as described (25Sargueil B. Pecchia D.B. Burke J.M. Biochemistry. 1995; 34: 7739-7748Google Scholar). The 3′ cleavage product was obtained by solid-phase synthesis. Ligation reactions were carried out with 10 nminternally labeled ribozymes (ribozyme-5′ cleavage product) and 10 μm 3′ cleavage product. Neither the rate nor the extent of ligation was changed by increasing the concentration of the 3′ cleavage product, indicating that 10 μm is enough to achieve saturation (data not shown). The fraction of ribozyme ligated was plotted versus time and fitted to the single-exponential equation shown above (Equation 1). The final extent of ligation was typically between 0.5 and 0.7. The equation parameters were fitted by means of nonlinear regression analysis as described above. Experimental and fitting errors were as described for the cleavage reaction. The 5′ cleavage product with a 2′,3′-cyclic phosphate was obtained from a preparativetrans-cleavage reaction using 5′-32P-substrate. The end-labeled 5′ cleavage product was then gel purified as described (25Sargueil B. Pecchia D.B. Burke J.M. Biochemistry. 1995; 34: 7739-7748Google Scholar). Ligation reactions were carried out with a small amount of 5′-32P 5′ cleavage product (less than 1 nm) and saturating excess of 3′ cleavage product and ribozyme (8 μm each). The fraction of 5′ cleavage product ligated to the 3′ cleavage product (substrate formation) was plottedversus time and fitted to the double-exponential equation shown above (Equation 2). The amplitudes and rates of the biphasic time course were estimated as described for the cleavage reaction. A small amount of 5′-32P-substrate (less than 1 nm) was incubated with a saturating excess of ribozyme (200 nm) in the reaction buffer for 2 min (wt ribozymes) or 30 s (SV5 ribozymes) at 25 °C. These incubation times allow essentially complete formation of the ribozyme·substrate complex since the binding half-times under these conditions are about 30 and 1 s, for the wt and the SV5 ribozymes, respectively (see “Results”). The chase step is initiated by adding a large excess (5 μm final concentration) of either a DNA oligonucleotide that is fully complementary to the wt substrate or, alternatively, a nonradiolabeled SV5 substrate for reactions carried out with wt or SV5 ribozymes, respectively. A complementary DNA oligonucleotide was used instead of the unlabeled wt RNA substrate because this molecule forms stable dimers at high concentrations. During the chase period, aliquots were removed and quenched with an equal volume of loading buffer (15 mm EDTA, 97% formamide). Samples were analyzed and quantified as described above. Parallel control reactions were carried out in the absence of the chase molecule. The efficiency of the chase step was evaluated by mixing the labeled substrate with the chase molecule (either the complementary DNA oligonucleotide or the unlabeled RNA substrate) prior to the addition of ribozyme. No significant cleavage of the labeled substrate was observed under these conditions, indicating that there is no rebinding of the labeled substrate during the chase step. Typically, time courses carried out in the presence of the chase molecule displayed monophasic behavior and, therefore, were fitted to single-exponential equations (Equation 1). Control reactions in the absence of chase showed biphasic kinetics, and hence double-exponential equations (Equation 2) were used. Estimation of the kinetic parameters (amplitudes and rates) was carried out as described above. Rate constants for substrate association were measured using a series of pulse-chase experiments, similar to those used to evaluate substrate dissociation. Several ribozyme concentrations, ranging from 12.5 nm to 200 nmfor wt ribozymes or from 1 nm to 10 nm SV5 ribozymes, were combined with a trace amount (less than 0.1 nm) of the corresponding 5′-32P-substrate in reaction buffer at 25 °C. For each ribozyme concentration, several chase reactions were initiated at different times, ranging from 10 s to 4 min. The chase molecule was a complementary DNA oligonucleotide, in the case of the wt substrate, or unlabeled RNA substrate, in the case of the SV5 substrate. The final concentration of the chase molecule was 5 μm or 1 μm for reactions carried out with wt or SV5 substrates, respectively. Reactions were incubated for 1 h at 25 °C after addition of the chase molecule. This time is sufficient to ensure a quantitative cleavage of the 5′-32P-substrate·ribozyme complexes (the half-time for the cleavage reaction is about 5 min). Time courses of the cleavage reaction were fitted to single-exponential equations, and the observed rates were plotted versus ribozyme concentration to obtain association (kon) and dissociation (koff) rates (2Herschlag D. Cech T.R. Biochemistry. 1990; 29: 10159-10171Google Scholar). Using the four ribozymes described above (wt, wt EH4, SV5, and SV5 EH4 ribozymes; Fig. 1) with corresponding substrates, we have carried out pre-steady-state and steady-state kinetic analyses. Pre-steady-state kinetics were used to measure individual rates for substrate binding (association and dissociation) and catalysis (cleavage and ligation). Steady-state analysis was used to assess rate-limiting steps and to estimate the interconversion rates between different conformations of the ribozyme (see below). Experiments in which ribozyme was in large excess over substrate (single-turnover conditions) were used to measure the first-order rate for cleavage of substrate (see “Materials and Methods”). Under these conditions, the observed rate will reflect the rate of the cleavage step, unless both cleavage products remain bound to the ribozyme long enough as to be ligated (reverse reaction). However, product dissociation is much faster than ligation in the hairpin ribozyme (14Hegg L.A. Fedor M.J. Biochemistry. 1995; 34: 15813-15828Google Scholar), preventing the interference of the ligation activity in a cleavage assay carried out in the presence of ribozyme excess. Time courses for cleavage catalyzed by wt, wt EH4, SV5, and SV5 EH4 ribozymes were followed at 25 °C. An excess of 200 nmribozyme was used with picomolar concentrations of their corresponding [γ-32P]ATP radiolabeled substrates to monitor cleavage rates under single-turnover conditions (see “Materials and Methods”). A typical time course for such a cleavage reaction, as catalyzed by the wt ribozymes is shown in Fig. 2. For all four ribozymes, the experimental data were fitted to single- and double-exponential equations (see “Materials and Methods”). As shown in Fig. 2, the data are much better described as biphasic rather than monophasic reactions. This was evident by visual inspection of the data and from the comparison of statistical error parameters (standard deviation, χ2, and determination coefficient,R 2). Table I lists the values of the rates and amplitudes for the two different reaction phases for all four ribozymes. For the rest of this paper, we will refer to these phases as “fast” and “slow” according to their corresponding rates. The amplitudes and rates of both the phases were independent of ribozyme concentrations (25–200 nm) under single turnover conditions (Fig. 3). This rules out the possibility that the biphasic behavior of the ribozymes is due to the aggregation of two or more molecules. Before initiation of the reaction, both the ribozyme and the radiolabeled substrate were incubated separately at 37 °C for 10 min. To induce different folding conditions, different preincubation protocols were followed before the cleavage reactions were initiated. These conditions included preincubation at 90 °C for 1 min, 65 °C for 10 min, 37 °C for 10 min, and no preincubation. In all cases, biphasic kinetics were observed.Table IKinetic parameters for the cleavage reactionRibozyme1-bRibozyme nomenclature is as defined in the Introduction and Fig. 1Fast phase1-aFast and slow phases correspond to the two components of the biphasic time courses.Slow phase1-aFast and slow phases correspond to the two components of the biphasic time courses.Amplitude1-cAmplitudes were normalized to the final extent of cleavage.Rate, min−1Amplitude1-cAmplitudes were normalized to the final extent of cleavage.Rate, min−1wt Rz0.540.050.460.005wt EH4 Rz0.760.330.240.02SV5 Rz0.740.160.260.01SV5 EH4 Rz0.770.110.230.005Cleavage reactions were carried out under single-turnover conditions, as described under “Materials and Methods.” Amplitudes and rates were obtained by fitting the experimental data to double-exponential equations (see “Materials and Methods”). Standard deviations from independent experiments were typically less than 50% for the parameters of the wt ribozymes and less than 25% for those of the SV5 ribozymes.1-a Fast and slow phases correspond to the two components of the biphasic time courses.1-b Ribozyme nomenclature is as defined in the Introduction and Fig. 11-c Amplitudes were normalized to the final extent of cleavage. Open table in a new tab Cleavage reactions were carried out under single-turnover conditions, as described under “Materials and Methods.” Amplitudes and rates were obtained by fitting the experimental data to double-exponential equations (see “Materials and Methods”). Standard deviations from independent experiments were typically less than 50% for the parameters of the wt ribozymes and less than 25% for those of the SV5 ribozymes. Except in the case of the wt ribozyme, approximately 75% of the substrate was cleaved in the fast phase of the reaction. The rate of the fast phase was generally at least 10-fold greater than the rate of the slow phase. Although the reaction rates follow a similar trend for the wt ribozyme, the amplitudes of the fast and slow phases were almost equal. This indicates that the partitioning of the two phases in the case of the wt ribozyme is different from the other three variants of the hairpin ribozyme. The rates of ligation for all four constructs of the hairpin ribozyme were measured using two different approaches, cis- and trans-ligation.Cis-ligation was carried out using a self-cleaving version of the hairpin ribozyme that is covalently attached to the 5′-product through a short pentacytidine linker (28Berzal-Herranz A. Joseph S Burke J.M. Genes Dev. 1992; 6: 129-134Google Scholar), as described under “Materials and Methods.” The cis-ligation rate was measured using a trace quantity of the internally labeled ribozyme (10 nm) with a saturating excess of 3′-product (10 μm). In contrast, the trans-ligation reaction was monitored using a trace amount of 5′-32P 5′-product (<1 nm) in the presence of saturating concentrations of 8 μm ribozyme and 3′-product (see “Materials and Methods”). In both cases (cis- andtrans-reactions), the observed ligation rate will reflect an approach to the equilibrium between cleavage and ligation since cleavage is much faster than dissociation of the ribozyme·substrate complex (see below). Therefore, the observed rate will be the sum of cleavage and ligation rates. Cis-ligation kinetics were studied for each of the four ribozymes, and reaction profiles followed single-exponential kinetics (Fig. 4 A and Table II). On the other hand, the ligation reaction in trans catalyzed by SV5 and SV5 EH4 ribozymes clearly showed biphasic kinetics (Fig.4 B and Table II). These ligation time courses were fitted to double-exponential equations, which allowed estimation of amplitudes and rates of both phases (Table II). Rates of trans-ligation for wt and wt EH4 ribozymes could not be measured since the high ribozyme concentration required for these reactions led to accumulation of ribozyme dimers (23Butcher S.E. Burke J.M. Biochemistry. 1994; 33: 992-999Google Scholar).Table IIKinetic parameters for the ligation reactionRibozyme2-dRibozyme nomenclature is as defined in the Introduction and Fig. 1.Cis-reactions2-aCis-ligation reactions were carried out as described under “Materials and Methods,” using a trace amount of internally labeled ribozyme, covalently attached to the 5′ cleavage product through a pentacytidine linker, in the presence of saturating excess of the 3′ cleavage product. Reactions carried out under these conditions displayed monophasic behavior. Rates were estimated by fitting the experimental data to single-exponential equations (see “Materials and Methods”).Trans-reactions2-bTrans-ligation reactions were carried out with a trace amount of end-labeled 5′ cleavage product, in the presence of saturating excess of both 3′ cleavage product and ribozyme. Reactions conducted under these conditions showed biphasic behavior.Rate, min−1Fast phase2-cFast and slow phases represent the two components of the biphasic time courses. Amplitudes and rates were obtained by fitting the experimental data to double-exponential equations (see “Materials and Methods”). Standard deviations from independent experiments were typically less than 25%.Slow phase2-cFast and slow phases represent the two components of the biphasic time courses. Amplitudes and rates were obtained by fitting the experimental data to double-exponential equations (see “Materials and Methods”). Standard deviations from independent experiments were typically less than 25%.Amplitude2-eAmplitudes were normalized to the final extent of ligation.Rate, min−1Amplitude2-eAmplitudes were normalized to the final extent of ligation.Rate, min−1wt Rz1.9wt EH4 Rz5.5SV5 Rz0.40.91.30.10.1SV5 EH4 Rz0.40.92.30.10.22-a Cis-ligation reactions were carried out as described under “Materials and Methods,” using a trace amount of internally labeled ribozyme, covalently attached to the 5′ cleavage product through a pentacytidine linker, in the presence of saturating excess of the 3′ cleavage product. Reactions carried out under these conditions displayed monophasic behavior. Rates were estimated by fitting the experimental data to single-exponential equations (see “Materials and Methods”).2-b Trans-ligation reactions were carried out with a trace amount of end-labeled 5′ cleavage product, in the presence of saturating excess of both 3′ cleavage product and ribozyme. Reactions conducted under these conditions showed biphasic behavior.2-c Fast and slow phases represent the two components of the biphasic time courses. Amplitudes and rates were obtained by fitting the experimental data to double-exponential equations (see “Materials and Methods”). Standard deviations from independent experiments were typically less than 25%.2-d Ribozyme nomenclature is as defined in the Introduction and Fig. 1.2-e Amplitudes were normalized to the final extent of ligation. Open table in a new tab The observed ligation rates were about 15-fold faster than the cleavage rates (compare Table I with II). Therefore, the rates shown in Table IIessentially reflect the ligation step since the reversal reaction (cleavage) is negligible. This indicated that the hairpin ribozyme is an efficient ligase. A similar observation was reported from a previous kinetic study with the hairpin ribozyme (14Hegg L.A. Fedor M.J. Biochemistry. 1995; 34: 15813-15828Google Scholar). Ligation rat" @default.
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• W2023363067 title "Kinetic Mechanism of the Hairpin Ribozyme" @default.
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