Group II introns: highly specific endonucleases with modular structures and diverse catalytic functions

Methods 28 (2002) 323–335 www.academicpress.com

Group II introns: highly specific endonucleases with modular structures and diverse catalytic functions Olga Fedorova,a,b Linhui Julie Su,a and Anna Marie Pylea,b,* a

Department of Biochemistry and Molecular Biophysics, 630 West 168 Street, Box 36, Columbia University, New York, NY 10032, USA b Howard Hughes Medical Institute, Columbia University, New York, NY 10032, USA Accepted 30 July 2002

Abstract Group II introns are large catalytic RNAs with a remarkable repertoire of reactions. Here we present construct designs and protocols that were used to develop a set of kinetic frameworks for studying the structure and reaction mechanisms of group II introns and ribozymes derived from them. In addition, we discuss application of these systems to structure/function analysis of the ai5c group II intron. Ó 2002 Elsevier Science (USA). All rights reserved. Keywords: Group II intron; Kinetics; Self-splicing; Ribozyme

1. Introduction Group II introns are autocatalytic RNAs found in the organellar genes of plants, algae, fungi, and yeasts and in bacterial genomes [1–3]. Removal of these introns is critical for the expression of housekeeping genes in many of these organisms [1]. Although some group II introns can self-splice in vitro, most require protein cofactors for splicing in vivo [4,5]. Self-splicing of group II introns occurs via two competitive pathways, both in vitro and in vivo [6,7]. In the transesterification pathway, the 20 -OH group of the branch-point adenosine serves as a nucleophile, which attacks the 50 -splice site and results in the formation of a branched lariat structure (Fig. 1A). This reaction is mechanistically similar to that of nuclear mRNA splicing by the spliceosome [8,9]. Alternatively, water can serve as a nucleophile for hydrolytic splicing, which results in the formation of a linear intron. This pathway is mechanistically similar to that of RNase P, and it is required for the splicing of group II intron families that do not contain branch-point structures [10,11]. In both

*

Corresponding author. Fax: 1-212-305-1257. E-mail address: [email protected] (A.M. Pyle).

pathways, the free 30 -OH of the 50 -exon serves as the nucleophile during the second step of splicing, leading to formation of ligated exons. Hydrolytic reopening of the exons can also be observed under certain in vitro conditions (spliced exon reopening; Fig. 1A; [12]). In addition to reactions involved in self-splicing, free group II intron lariats can form RNP particles with their encoded maturase proteins [4,13]. These particles can behave as transposable elements that insert themselves into DNA and RNA targets (Fig. 1A) [14,15]. Despite the diversity of their reactions, the kinetic behavior of group II intron ribozymes is often remarkably simple, making them good model systems for studying mechanisms of RNA catalysis. The secondary structure of group II introns is highly conserved [1,2] and consists of six domains (Fig. 1B). The largest of these, domain 1 (D1), recognizes 50 exonic sequences, contains certain active-site elements [16,17], and may serve as a scaffold for binding of other domains [18]. Domain 2 (D2) contains certain long-range tertiary interactions [19,20] that may facilitate splicing. Domain 3 (D3) acts as a cofactor that greatly enhances the chemical rate of catalysis by group II introns [21,22]. Domain 4 (D4) contains an open reading frame (ORF) in certain group II introns. This ORF encodes a unique protein cofactor (the ‘‘maturase’’) that generally

1046-2023/02/$ - see front matter Ó 2002 Elsevier Science (USA). All rights reserved. PII: S 1 0 4 6 - 2 0 2 3 ( 0 2 ) 0 0 2 3 9 - 6

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Fig. 1. (A) Reactions catalyzed by a self-splicing group II intron. (Top) In hydrolysis pathway, water is the nucleophile attacking the 50 -splice site, resulting in formation of linearized intron and ligated exons. (Bottom) In branching pathway, the 20 -OH of a bulged adenosine in D6 attacks the 50 splice site, resulting in formation of a lariat intron and ligated exons. In the presence of a self-encoded protein, the lariat can insert itself into a target RNA or DNA. (B) Schematic diagram of the secondary structure of group II intron. Shaded boxes represent base-pairing sites between the 50 exon and the intron. The intron can catalyze hydrolysis of a short oligonucleotide RNA that mimicks the 50 splice site (bottom).

contains RNA recognition, reverse transcriptase, and DNA endonuclease motifs that assist in both intron splicing and mobility [2,23,24]. Domain 5 (D5) is the most conserved feature of the intron and, along with D1, it is absolutely required for all forms of catalytic activity [25–27]. Domain 6 (D6) is not highly conserved, yet it supplies the bulged adenosine that is essential for the branching pathway of splicing (Fig. 1A). Active forms of a group II intron can be assembled from individual domains that are transcribed separately. This remarkable characteristic has facilitated the design of ribozyme constructs that contain selected domains, thereby allowing investigators to study the role of individual intron substructures in the reaction mechanism. Optimized kinetic frameworks have been developed for many of these ribozyme constructs, and these have facilitated a diversity of functional and structural studies. Here, we present protocols for the kinetic analysis of ribozymes derived from the ai5c group II intron. The assays provided here are applicable for studying other self-splicing introns, other forms of catalytic RNA, and even complex assemblies of RNA and protein enzymes.

analysis provides the basis for designing meaningful future structure–function studies. Many enzyme–substrate systems are very complex, with different products formed along the reaction pathway. Adding to the complexity, these products may be formed by sequential or parallel mechanisms and some reaction steps may be fully or partially reversible. However, these features do not necessarily make an enzyme difficult to study. The ai5c group II intron is a good example of a ribozyme that is kinetically well characterized despite many of the complex issues described above. In vitro self-splicing of this intron results in the formation of several species: lariat intron, spliced exons, linear intron, and free 30 and 50 exons (Fig. 2A) [6,28,29]. There are two major approaches to studying the kinetics of the group II ribozyme-catalyzed reactions. The first approach involves monitoring complete self-splicing of a construct that contains all intronic domains and flanking exons. The second approach dissects the intron into modified constructs that allow one to focus on only one reaction. Each approach has advantages and disadvantages. Therefore, it is recommended to use a combination of both for characterizing a system of interest.

2. Kinetic analysis of a large, multifunctional ribozyme

2.1. Kinetic assay of the self-splicing ribozyme construct

To understand the mechanism of any process catalyzed by a protein or an RNA enzyme, one must perform a detailed kinetic analysis on the system. Such

There are certain advantages in performing kinetic studies on self-splicing of the intact intron. This approach provides information about accumulation of

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Fig. 2. (A) Comparison of ai5c self-splicing products and reactions. The bands are assigned as follows: A, lariat intron; B, precursor; C, linear intron; D, spliced exons; E, free 30 exon; F, free 50 exon. Time points were chosen to show similar amounts of reaction under different conditions. Lane 1 is the precursor RNA at t ¼ 0. For lanes 2–20, the reaction times are 5, 20, 65, 900, 1.5, 20, 45, 6, 14, 50, 30, 70, 160, 35, 75, 170, 30, 70, and 160 min., respectively, under the reaction conditions shown. Lane M contains RNA markers of the molecular masses shown. (B) Reaction profile in 100 mM MgCl2 and 0.5 M NH4 Cl. Reprinted with permission from JMB.

each splicing product, the existence of any intermediates, and the functional relationship between the individual reactions involved in splicing and the effect of

reaction conditions [29,30]. Mutational analysis in the context of the intact intron allows one to approximate the dynamic interplay of individual active-site elements

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along the splicing pathway. For these reasons, the kinetics of self-splicing has been intensively studied for the ai5c group II intron [6,28,29]. In one set of studies, the ai5c group II intron and its flanking exons was transcribed from the HindIII-linearized plasmid pJD20 [6,29]. Transcription is carried out in the presence of [a-32 P]UTP to obtain an internally labeled precursor RNA. The RNA is gel-purified and used in kinetic assays at 1 nM final concentration. The concentration of the radiolabeled RNA is determined based on the specific activity of [a-32 P]UTP. The splicing reactions are performed at 42 °C in 40 mM 30 -Morpholinopropanesulfonic acid (Mops), pH 7.5. The reaction buffer is supplemented with one of the following combinations of salts: 10 mM MgCl2 and 5 nM protamine; 100 mM MgCl2 ; 100 mM MgCl2 and 0.5 M KCl; 100 mM MgCl2 and 0.5 M NH4 Cl; or 100 mM MgCl2 and 0.5 M (NH4 )2 SO4 [6]. Here, splicing buffers without any monovalent salts are referred to as ‘‘low-salt’’ buffers; those containing potassium or ammonium salts in addition to magnesium chloride are ‘‘high-salt’’ buffers. RNA is first denatured in the presence of Mops at 95 °C for 1 min to eliminate misfolded conformations and allowed to cool to the reaction temperature by incubating the sample at room temperature for 1 min; then the salts are added and the resulting mix is incubated at 42 °C. Aliquots are removed at different time points, quenched with an equal volume of gel-loading buffer (1 TBE, 1.8% sucrose, 0.02% xylene cyanol, 36% (v/v) formamide, and 25 mM EDTA), and analyzed on a 4% denaturing gel. Linear and lariat intronic species were observed under all conditions (Fig. 2A). However, lariat intron species are more abundant in the presence of ammonium salts, whereas potassium chloride promotes formation of the linear intron. In ‘‘low-salt’’ buffers bands corresponding to spliced and free exons are sometimes obscured by the high background that is induced by random Mg2þ -induced RNA degradation [6]. During early studies of ai5c self-splicing, one of the most important questions was whether the linear intron is a product of lariat degradation or instead is formed independently through a competing hydrolytic pathway during the first step of splicing [28,29]. Any large RNA is susceptible to degradation, especially at late time points. A single cut anywhere in the lariat molecule would linearize it, resulting in species that comigrate with the linear intron on a denaturing gel. One feature that might distinguish truly linear intron from ‘‘broken lariat’’ is the absence of the branched structure. Isolation and nuclease mapping of linear species that accumulate at early time points did not yield any branched structures. However, at late time points (>2 h), the fraction of ‘‘broken lariat’’ among the linear products reached 45%. This strongly suggested that the linear intronic species is produced predominantly through a

hydrolytic pathway, although it becomes contaminated over time by degradation of the lariat. Additional corroborating evidence for a hydrolytic pathway came from two sources: (1) efficient splicing by mutant group II introns with deleted branch-points [7,31] and (2) detailed kinetic analysis of the wild-type intron, which demonstrated that accumulation of linear and lariat intronic products results from two competing pathways (branching and hydrolysis). Splicing reactions obey classic parallel kinetics [6,32] and cannot be fit using a sequential model in which the lariat serves as an intermediate for breakdown into linear species. Formation of the linear intron occurs at early time points and accumulates at a constant ratio with lariat products throughout the reaction pathway (even at later time points), indicating that linear intron is produced through an independent pathway that directly competes with branching. One feature that simplifies the kinetic analysis of ai5c self-splicing is that the first step is rate-limiting. No accumulation of lariat-30 exon intermediate is observed (Fig. 2A) [6,33]. Kinetic data that are specific for the first step of splicing can therefore be obtained by quantitating bands corresponding to precursor and products using radioanalytic imaging (Phosphorimager; Molecular Dynamics). To determine reaction rates in ‘‘low-salt’’ buffers, fractions of precursor and products are plotted versus time (t) and fit to simple single-exponential expressions, Precursor ðtÞ ¼ eks t Precursoro

ð1Þ

and Product ðtÞ ¼ 1  eks t ; Precursoro

ð2Þ

where ks is the total reaction rate. To calculate the rate constants for individual reactions (branching and hydrolysis) one multiplies ks by the fraction of the corresponding product. In ‘‘high-salt’’ buffers the accumulation of linear and lariat introns over time is biphasic and data can be fit using double-exponential equations. This behavior is thought to be caused by the presence of two populations of precursor molecules, A and B, that react at different rates [6] (Fig. 2B): Product ðtÞ ¼ fA ð1  eks t Þ þ fB ð1  eks t Þ: Precursoro

ð3Þ

The sensitivity of each pathway to ionic conditions is an important property of the group II ribozymes [30]. By varying the monovalent cation in the reaction mixture one can force the ribozyme to prefer one reaction pathway over the other. In ‘‘low-salt’’ and KClcontaining buffers the linear intron is the predominant product over the lariat, while in the presence of am-

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monium sulfate the formation of the lariat intron is favored over linear species [6,28,29]. When the splicing buffer is supplemented with ammonium chloride, both intronic products are formed in equal quantities [6].

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an alternative pathway is observed only in the presence of 1 M KCl.

3. Two-piece ribozyme constructs based on group II introns

2.2. Applications Determining the group II intron elements critical for activity. In vitro kinetic assays were used to assess the role of different intronic domains in the group II splicing [34]. A series of deletion constructs lacking individual or multiple intronic domains was created and tested for self-splicing activity. The use of a complete splicing assay in these studies has made it possible to approximate the importance of intronic domains for different splicing reactions. The same approach was used to determine the three catalytically important nucleotides in D5 (A816, G817, and C818), which are referred to as the ‘‘catalytic triad’’ [35]. Kinetic studies of point mutants in the context of an intact intron have unambigously demonstrated that the ‘‘catalytic triad’’ is essential for both pathways of self-splicing. Branch-point recognition studies. The kinetic approach described above has also been successfully applied for identifying determinants of group II branch-point recognition, using a series of intronic constructs with base and single-atom mutations in the vicinity of the branch-point [36,37]. The results of these studies provided a comprehensive model of the branch-point selection in group II intron splicing. Characterization of alternative splicing pathways. Although hydrolysis is not the dominant mechanism for splicing of most introns, it is a viable default pathway for ai5c mutants that contain severe branching defects both in vitro and in vivo [7]. Thus, hydrolysis is an important alternative to branching and it is not an artifact caused by nonphysiological reaction conditions. It is notable that an entire family of group II introns has now been found to lack a branch-point adenosine in domain 6. These introns self-splice through the hydrolytic pathway [11]. Comparison of diverse introns. Most group II introns require similar ionic conditions (high concentrations of magnesium and monovalent ions) to support self-splicing activity [29,38,39]. Interestingly, a group II intron from the brown algae Pylaiella littoralis (PL.LSU 2) has been reported to be very active under unusually low Mg2þ concentration (10 mM). However, this intron maintains a requirement for relatively high concentrations of monovalent salts (1 M) [40]. Remarkably, this intron does not splice via a hydrolytic pathway in the presence of ammonium salts, and trace amounts of linear species are attributable to the presence of broken lariat. Hydrolysis as

Despite the advantages of using full-length selfsplicing constructs as discussed above, interpreting data from self-splicing kinetics remains relatively complicated. Such complexity hinders complete kinetic and mechanistic characterization of individual group II intron reactions. Furthermore, different reactions do not necessarily involve the same elements of the ribozyme active site; conformational rearrangements are thought to occur along the splicing pathway [19,41,42]. For these reasons it is often more convenient to convert the whole intron into a ribozyme–substrate system that catalyzes only one reaction. The use of such systems allows one to measure binding and rate constants for each step of splicing separately and to control the chemical composition and concentration of individual domains. 3.1. A two-piece system to study the first step of splicing via hydrolysis Although multipiece constructs have been used to study D1, D3, and D5, the latter has been intensively analyzed using this approach. D5 can be transcribed or synthesized chemically as a separate molecule and added to other domains to promote group II reactions [26,27,43]. The first two-piece ribozyme system based on a group II intron involved addition of D5 to exD123 [26]. D5 catalyzes hydrolysis of the 50 exon from the intronic component. In this setup, exD123 undergoes a chemical transformation and is therefore treated as the reaction substrate. D5 remains unchanged in the course of the reaction and is treated as an enzyme. The stereochemistry of exD123 hydrolysis by D5 is equivalent to the first step of splicing via hydrolysis [12]. This finding, together with similar mutational sensitivities and rate constants to full-splicing systems, suggests that the exD123/D5 system is a good mimic for the first step of splicing. A detailed kinetic framework for this system was developed [44] and used to determine interdomain affinities and the chemical rate constant for D5-catalyzed hydrolysis. Single-turnover (STO) kinetic assay. Single-turnover kinetic assays have been carried out to characterize the kinetic behavior of product evolution with respect to time. In this experimental setup, the enzyme is provided in excess and the substrate in trace amounts. As such, neither substrate binding nor product off-rate will be rate limiting and the reaction rate should reflect either chemistry or a conformational change within the enzyme–substrate complex. STO reactions are carried out

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using internally labeled exD123 at 1 nM and unlabeled D5 in a range of concentrations from 50 nM to 6 lM. Reactions are performed in a total volume of 10–20 ll in a buffer containing 40 mM Mops, pH 7.5, 100 mM MgCl2 , and 0.5 M KCl. The ribozyme and substrate are heated separately at 95 °C for 1 min to denature any misfolded structures and allowed to cool to the reaction temperature of 45 °C. The reaction is initiated by combining the RNA molecules with a simultaneous addition of MgCl2 and KCl. Aliquots (0.5–1 ll) are removed throughout a 60-min time course and quenched with formamide gel loading buffer (see above). The samples are then analyzed on a 5% denaturing gel. Apparent rate constants (kobs ) are calculated from the slope of a semilog plot [ln(fraction unreacted exD123) vs time]. To test whether this pseudo first-order treatment is applicable, the concentration of labeled exD123 (the component in trace) is varied within an order of magnitude (0.3 to 3 nM). A constant kobs in this range suggests proper pseudo first-order behavior. In addition, a semilog plot of the fraction of unreacted precursor (exD123) versus time should remain linear for three to five halftimes [45]. To determine kinetic parameters kcat (ST) and KM (ST), reactions are carried out at different concentrations of D5. Apparent rate constants (kobs ) are then plotted against [D5] and fit to a simple 1:1 binding isotherm (Fig. 3):
kobs ¼ kcat ðSTÞ ½exD123 t þ ½D5 t þ KM

qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi  ð½exD123 t þ ½D5 t þ KM Þ2  4  ½D5 t  ½exD123 t  ð4Þ ð2  ½exD123 t Þ:

Because D5 is not allowed to turn over, product release can be rate-limiting and kcat (ST) from the plot represents the maximum rate of the process. To deter-

Fig. 3. Determination of KM (ST) and kcat (ST) by plotting kobs values (inset) as a function of [D5]. Kinetic constants were determined from the fit of experimental data to Eq. (4). Reprinted with permission from Biochemistry.

mine whether kcat (ST) and KM (ST) represent the rate of the chemical step and a true binding constant, STO kinetics should be complemented with other experiments described below. Direct binding assay. To determine whether KM represents the true binding constant (Kd ), D5 binding to exD123 was measured directly. In this experiment, D5 is 50 -end labeled and in trace (1 nM final) and exD123 is unlabeled and in excess at varying concentrations from 50 nM to 5 lM. D5 and exD123 are denatured, combined, and supplemented with salts as described in the STO kinetic protocol. The samples are then incubated at the reaction temperature for 15 min to ensure complex formation. Bound D5 is separated from unbound D5 by gel filtration. The procedure is carried out either on a long column packed with Sephacryl S-100 equilibrated with the reaction buffer, or on spin columns packed with the same matrix. In the first case, the sample is loaded onto the 8-ml column directly after the incubation and eluted with a flow rate of 1 ml/min. In this setup the bound complex is eluted right after the void volume. To determine the amount of unbound D5, fractions (100 ll) are collected during elution and placed into scintillation vials for Cerenkov counting. To determine Kd , the fraction of bound D5 is calculated and plotted versus exD123 concentration and the data are fit to a simple 1:1 binding isotherm [46,47]:
h ¼ ½D5 tot þ½exD123 tot þKd qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi  ð½D5 tot þ½exD123 tot þKd Þ2 4½D5tot ½exD123 tot  ð2½D5 tot Þ ð5Þ This method has been shown to overestimate Kd , presumably due to the lengthy procedure [44]. Therefore, the use of spin columns for separation of bound and unbound D5 has been developed as an alternative approach [48,49]. In this case empty spin columns are packed with 600 ll of Sephacryl S-100 equilibrated as described above. The sample is loaded onto the column and spun for 15 s, after which 25 ll of the buffer is added to the column and the sample is spun again for 30 s. Gel matrix is then separated from the eluate and 32 P-labeled D5 is quantitated using a scintillation counter. Data analysis is performed as described above. Kd is determined from the fit and compared to a KM (ST) value estimated from STO. It is important to note that the spin column gel filtration protocol provided here was specifically optimized for the separation of free D5 (34 nt) from D5 bound to exD123 (1003 nt). When working with molecules of other sizes, one would need to vary the volume of elution buffer, spinning time, and sometimes the matrix to achieve optimal separation (see [50] for an example).

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Multiple-turnover (MTO) kinetic assay. Multipleturnover kinetics experiments provide additional mechanistic information and may validate kinetic parameters obtained under single-turnover conditions. Furthermore, since enzyme is the limiting component in a multiple-turnover reaction, the experiment can be designed to provide an estimate of the active enzyme population. In contrast to STO reactions, the substrate is in excess and the ribozyme is in trace during MTO experiments. To visualize products, a trace amount of internally labeled exD123 is added to an excess of unlabeled exD123 (20–250 nM). D5 is added at low concentrations (5 nM) to ensure at least fourfold excess of the substrate at all times. Ideally, the concentration of D5 in these experiments is much lower than Kd . Reactions are initiated the same way as before, but a longer time course is needed to detect products. MTO studies of exD123 cleavage are significantly more difficult to perform than STO studies, as they require longer incubation times that promote Mg2þ -induced degradation of exD123, resulting in a high background. Initial rates of hydrolysis (kobs ) are calculated for each time course and then represented on an Eadie–Hofstee plot. In this formalism, kobs is plotted versus kobs /[S] to determine KM and kcat [44]. For D5-catalyzed hydrolysis of exD123, KM values from single- and multiple-turnover experiments and the directly measured Kd value are in a good agreement [44]. These results suggest that the majority of the D5 and exD123 populations are functionally active, and the rate of catalysis is unlikely to be limited by a conformational change within the D5–exD123 complex. However, to definitively test whether kcat ¼ kchem , it is usually advisable to conduct additional tests on the sensitivity of kcat to pH and/or to single-atom changes at the site of reaction. Mechanistic evaluation of the kcat from single turnover. If kcat from STO experiments represents the chemical step of the reaction catalyzed via a general acid–base mechanism, this value will be limited by the rate of proton transfer and it will depend on the pH of the reaction buffer. To determine whether this was the case for the exD123–D5 system, reaction rates at saturating [D5] were measured at different pH values varying from pH 5 to 9. To sample this broad pH range, different buffers are used. However, it is important for the pH ranges of the buffers to overlap to ensure that reaction rates remains the same in different buffers at the same pH. The data from the time courses are plotted as log(kobs ) vs pH. If the fit is linear, with a slope close to 1, then the rate is likely to represent a chemical step in which proton transfer is rate-limiting. For D5-catalyzed hydrolysis of exD123, the rate of catalysis was found to be limited by chemistry in the pH range from 5 to 7. This behavior suggests that titratable groups with pKa values close to 7 are involved in general base catalysis at the ribozyme active site. Similar tests for rate-limiting chemistry are

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provided by incorporating phosphorothioates in the substrate and examining rate effects [51]. Application of D5–exD123 system to structure–function studies. All of the above experiments suggest that the D5-catalyzed hydrolysis of exD123 obeys simple Michaelis–Menten kinetics and that, under single-turnover conditions, the apparent kinetic parameters KM and kcat represent binding and chemical rate constants, respectively. Given this kinetic mechanism, it was possible to study the effect of different mutations and single-atom changes in D5 and attribute them to specific roles in the ground state or transition state of reaction [49,52,55]. Extensive phylogenetic analysis of group II introns suggested that there is no possibility for D5 to form Watson–Crick base pairs with exD123 [1,2]. Therefore, binding was presumed to occur via long-range tertiary interactions involving participation of the sugar–phosphate backbone. Because a D5 molecule consisting entirely of DNA does not support either exD123 binding or chemistry [53], at least some of 20 -OH groups were likely to be important for ground state and transition state interactions. To elucidate the mechanistic contribution of individual 20 -OH groups, a series of D5 molecules containing single 20 -deoxy substitutions was chemically synthesized and tested for catalytic activity during STO reactions with exD123 [53]. These experiments identified D5 positions in which 20 -deoxynucleotide substitution resulted in >10-fold effects on either kcat or KM , suggesting that the corresponding 20 -OH groups play important and highly specific roles in the reaction mechanism. The same approach was used to examine the importance of functional groups on the most highly conserved nucleotide of D5 (G817). This guanosine is part of the catalytic triad and it is among the most critical elements in the intron active site [35,52]. D5 molecules containing single-atom changes at G817 were synthesized and reacted with exD123 under STO and MTO conditions [49]. It was found that major groove substitutions at O6 and N7 of G817 severely inhibited reaction chemistry but not binding, as demonstrated by competition and direct binding assays. By contrast, minor groove substitutions at N2 affected binding, but not the chemical rate. These findings elucidated the mechanistic contributions of G817 and, in a more global sense, helped to define the specific roles of the D5 major and minor grooves in transition state stabilization and D5 docking, respectively [49,53]. In addition to the powerful enzymatic assays provided by the D5–exD123 system, the strong complex between these molecules facilitated chemical footprinting studies for analysis of tertiary contacts between D5 and D123 [50]. This approach is based on the fact that exD123 functional groups will have differential susceptibility to attack by dimethyl sulfate (DMS) in the

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Fig. 4. Time course of branching independent of hydrolysis. (A) PAGE showing 50 -end-labeled D56 (1 nM) reacting with cold exD123 (2 lM), which results in formation of D56D123. (B) Quantification of product evolution plotted with respect to time. Reprinted with permission from RNA.

presence and absence of D5 [54,55]. DMS footprinting studies elucidated exD123 sites that represent thermodynamically significant contacts between D5 and exD123. 3.2. Two-piece system for studying the first step of splicing via branching A different two-piece construct was designed to examine the first step of splicing via branching. To study this pathway, an RNA consisting of domains 5 and 6 (D56) is transcribed separately and reacted with exD123 in trans [56]. To prevent the second step of splicing, the D56 RNA transcript does not contain any nucleotides beyond the 30 splice site. A competing hydrolytic pathway also takes place in this system. However, the assay can be set up in a way that branching is monitored independent of hydrolysis, allowing one to focus solely on the kinetic parameters describing transesterification. Studying branching independently of hydrolysis. The experimental setup for studying the D56 reaction with exD123 is similar to that described previously for the two-piece hydrolysis system (40 mM Mops, pH 7.0, 100 mM MgCl2 ), except that 0.5 M NH4 Cl is used instead of KCl to specifically promote branching. When D56 reacts with exD123, the bulged adenosine in D6 attacks the 50 splice site in exD123, releasing the 50 exon and resulting in a Y-shaped product that contains a 20 –30 –50 branched nucleotide that joins D56 to the first nucleotide of the intron (in D123). To monitor the branching reaction independent of hydrolysis, 50 -endlabeled D56 (in trace) is reacted with excess exD123 (Fig. 4A). In this type of trans-branching assay, the reaction fits a perfect single exponential, although the reaction amplitude never exceeds 50% (Fig. 4B) [56]. By

varying the concentration of exD123 used in excess, one can determine the dependence of kobs on [exD123], which is fit to a standard bimolecular 1:1 binding equation as previously described. Remarkably, Kd from the fit is equal to Kd for the hydrolysis system (exD123– D5), implying that D6 does not contribute to binding and is presented to the active site by D5. Assay to test the reversibility of branching. When a reaction cannot exceed a certain low amplitude, as in the case of D56 branching, there are at least two likely explanations: (1) a large fraction of D56 is inactive and (2) the reaction is reversible. The former can be ruled out in a number of ways, including isolation of ‘‘unreacted material’’ and testing to determine whether it can react. The latter can be assessed by running the reaction in reverse. To test reversibility of D56 transesterification, the normal reaction products (D123D56 and 50 exon) are isolated and used as starting materials. One then monitors the formation of precursor molecules (exD123 and D56). For this purpose, labeled D123D56 obtained via trans-branching (1 nM) is isolated and reacted with an excess of free 50 exon (>10 nM, a concentration that is higher than Kd reported for the 50 exon/intron binding [57]) under reaction conditions that are similar to those used in the branching reaction described above. The release of D56 from D123D56 in the presence of 50 exon (which attacks via its 30 -OH nucleophile) is monitored over time and products are analyzed and quantitated as described previously. When the fraction of reacted D123D56 is plotted versus time, the kinetic curve is biphasic: a fast burst (first 20% of the product formed) is followed by a slow phase (Fig. 5). The curve can be fit with a double-exponential equation, y ¼ Aek1 t þ ð1  AÞek2 t , where A represents the fraction of molecules participating in a fast phase, k1 is the rate of the fast phase, and k2 is the rate of the slow phase. The

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Demonstrating that D123D56 can be converted to D56 in the presence of a 50 exon is not sufficient to conclude that branching is reversible. It is possible that 50 exon just accelerates the natural hydrolysis of 20 –50 linkage in D123D56. If branching is truly reversible, 50 exon should become covalently attached to D123 after D56 is released. To test this assumption, 50 -end-labeled 17-mer exon (2 nM) is reacted with 0.1 nM of the branched product (D123D56). The mobility shift of 50 labeled exon, corresponding to exD123 formation, indicates that branching is fully reversible.

4. Establishing a system to study ribozyme folding and substrate recognition via kinetics Fig. 5. Kinetic analysis of the reverse-branching reaction. Fraction of reacted D56D123 was plotted with respect to time and fit to a doubleexponential equation. Reprinted with permission from RNA.

apparent kinetic behavior is attributable to two populations of the precursor molecules: one in a favorable conformation and ready to react and the other in unfavorable conformation and in need for a conformational change to become active.

In all of the precursor constructs discussed above, the 50 exon is attached to the intron. To study 50 splice site recognition in more detail, it is useful to design a system in which the exonic component is synthesized separately and presented in trans to the intronic components (Fig. 6A) [58–60]. The 50 exonic substrate consists of IBS 1 and 2 and the first seven nucleotides of the intron (Fig. 6B). Systems with exonic substrate in trans have a wide variety of mechanistic and structural applications, in-

Fig. 6. (A) Scheme showing cleavage of a short RNA substrate (S) by a group II ribozyme. Domain 1 (D1) combines with domain 5 (D5) and substrate (S) to catalyze specific cleavage at the site analogous to the 50 splice site. Shaded regions and dashes between D1 and S designate pairing interactions IBS1–EBS1 and IBS2–EBS2. Reprinted with permission from Biochemistry. (B) Base-pairing scheme between EBS region of the substrate and BBS region of the ribozyme. Arrows denote positions of nucleotide substitution. Inverted triangle indicates the cleavage site. Reprinted with permission from JMB.

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cluding studies of folding, substrate specificity, and determinants for cleavage site selection. When designing the ribozyme component of the system, one can exploit the modular domains of a group II intron and create a variety of constructs. Cleavage of oligonucleotide substrates can be studied with a onepiece ribozyme (the D135 ribozyme, where the three domains are covalently attached in the proper sequence) or using a two-piece ribozyme construct (D1 or D13 with D5 provided as a separate molecule). Because reaction rates vary drastically from one system to another [0.6 min1 for D135 vs 0.003 min1 for D1 D5 (a dot represents a ribozyme system formed from two pieces) under the same reaction conditions (80 mM Mops, pH 7, 100 mM MgCl2 , 500 mM KCl)], an appropriate system should be chosen for studying a particular mechanistic or structural feature. 4.1. Ribozyme system for studying substrate specificity and fidelity For studying substrate specificity and fidelity, variants of a two-piece ribozyme system (D1 or D13 with D5 and oligonucleotide substrate in trans) were used. Comprehensive kinetic frameworks for both systems (both D1 and D13) were developed [57,61,62]. The experimental setup for kinetic analysis was similar to that previously described for the exD123 D5 system (see above). 4.2. Applications Determining the role of 20 -OH groups on the substrate backbone. The D1 D5 and D13 D15 ribozymes allow facile probing of functional groups critical for substrate hydrolysis. Specifically, it was of interest to determine whether the 20 -OH groups on the oligonucleotide substrate were important for the reaction. To address this question, single and block deoxynucleotide substitutions were incorporated along the oligonucleotide substrate in the IBS2, IBS1, linker, or the intronic region [61]. The slope of the pH–rate profile was 1 from 6.5–7.5 for the cleavage of deoxynucleotide substrates, suggesting that the reaction was limited in rate by chemistry [61]. Furthermore, deoxynucleotide substitution on the substrate resulted in only modest effect on KM (2–3) [61]. Together, these results suggest that the group II ribozymes cleaves DNA linkages almost as readily as RNA linkages [61]. These findings are consistent with the fact that group II introns are mobile genomic elements that can act as DNA endonucleases in the presence of self-encoded proteins [14,15,63]. A high-substrate specificity and a unique cleavage site selection for group II-derived ribozymes. The ability of group II introns to readily cleave DNA and RNA substrates and their role as mobile elements have engen-

dered interest in using group II intron ribozymes as a tool for genetic engineering [64–66]. As for any tool that may someday be used to manipulate genetic structure, it is of central importance to understand the substrate specificity of group II ribozymes and their propensity to cleave at the proper position. It is worth noting that the recognition site between the substrate and the intron is unusually long (13 bp (Fig. 6B)) and distributed between two EBS elements located far apart on the intron RNA [62]. Though the recognition sequences of group II introns (EBS1 and 2) are not conserved among group II members, their covariation in Watson–Crick basepairing with exon is phylogenetically conserved, indicating that the intronic recognition sequences can be redesigned to cleave a specific target sequence [1,63,67]. To quantitatively monitor target site specificity by a group II intron, and to determine how frequently it will choose the correct vs an incorrect sequence, the D13 D5 ribozyme was used to cleave matched and mismatched oligonucleotide substrates [62]. Single-nucleotide mutations were introduced along IBS regions of the exonic substrate, the specificity index (kcat =KM value) was measured for all mutant substrates, and these values were then compared to that of the wild-type substrate. It was found that while some mismatches led to a weaker KM , nearly all mismatches along the IBS region resulted in a significant drop of kcat . This suggests that specificity is maintained at two levels: a weaker KM suggests that the mismatched substrate has a faster koff . The drop in kcat implies that mismatched substrates have a greater likelihood of falling off the binding site uncleaved [62]. These results suggest that group II introns have remarkably high substrate specificity and tolerate mismatches poorly [62]. Given that group II intron-derived ribozymes have unusually high substrate specificity, it was of interest to determine how accurately they cleave a specific site once they are bound to a target sequence (cleavage fidelity). For example, even if a ribozyme always binds the correct target, it may not choose the site of cleavage with high accuracy. To evaluate this, single-nucleotide mutations were introduced downstream of the normal cleavage site (in the +1–+7 region outside the IBS sequences; Fig. 6B [68]). Interestingly, while most downstream mutations had little effect on cleavage rates or site selection [33,68,69], substitution of +1G to a C resulted in loss of fidelity [33,68]. This interesting observation led to a detailed kinetic analysis using substrates in which secondary mutations were introduced at various positions along the IBS1 sequence to determine whether the stability of the EBS–IBS helix was important for cleavage site selection. Based on these results, it appears that group II introns will cleave after the most thermodynamically stable basepairing between the ribozyme and its target substrate. The ribozyme core detects the terminus of the

O. Fedorova et al. / Methods 28 (2002) 323–335

EBS–IBS helix and cleaves at the junction between duplex and single-stranded regions [68].

5. Applications of kinetic assays to RNA folding Designing a folding construct. To create a suitable construct for folding studies, one should meet two criteria: the RNA construct should contain all components that are critical for function, and it must be conformationally homogeneous. For these reasons D135 is the most appropriate ribozyme construct. The ribozyme D135 includes all critical domains and is capable of only one reaction, the cleavage of a short oligonucleotide substrate in trans via hydrolysis [70]. Kinetics as a tool to probe for a homogeneous population of group II ribozyme. Because the D135 ribozyme can cleave substrate under multiple-turnover conditions, it is a useful probe for determining the fraction of the active ribozymes in a population. Active-site titration experiments are performed to determine reaction conditions in which the majority of the ribozyme population is catalytically active [71]. If the first turnover of reaction is fast and limited in rate by chemistry and subsequent turnovers are limited by the rate of product release, ‘‘burst’’ kinetics can be observed. Because the burst amplitude represents the amount of product formed in the first turnover, the magnitude of the burst represents the percentage of enzyme folded in the active conformation [70,71]. A crucial aspect of active-site titration experiments is the enzyme/substrate ratio. Typically, the oligonucleo-

Fig. 7. (A) Scheme for a kinetic assay for determining folding rate. (B) Product evolution is plotted with respect to time and fit to Eq. (6). Intercept of the short dashed line to the y-axis represents burst amplitude. Slope of the long dashed line represents burst rate. A burst rate constant of 1.0 min1 , with a subsequent turnover rate constant of 0.14 min1 (comparable to product release rate), was observed. Reprinted with permission from JMB.

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tide substrate is maintained in 10-fold excess with respect to the ribozyme concentration, in order to maintain a good signal/noise ratio while minimizing errors associated with experimental setup (for example, pipetting). In addition, we suggest making a substrate stock, containing trace-labeled substrate (for example, 1 nM labeled substrate combined with excess cold substrate to make a 200 nM substrate stock), to reduce background signals. The experiment is carried out using 4 nM D135, 40 nM substrate stock, 80 mM Mops, pH 7, 100 mM MgCl2 , and 500 mM KCl at 42 °C. Time points are taken at every minute during the first turnover and then every 5–10 min afterward. Data are analyzed as previously described. Equivalents of product is plotted against time and fit to a double-exponential equation, P ¼ A1 ð1  ek1 t Þ þ A2 ð1  ek2 t Þ;

ð6Þ

where P is equivalents of product, A1 is amplitude of the first turnover, k1 is burst rate (chemistry), A2 is amplitude of subsequent turnovers, and k2 is product release rate. Using burst titrations, we showed that >80% of D135 is active during all experiments. It is important to note that ideal reaction conditions may vary with each ribozyme construct. As such, one should consider varying salt, pH, and temperature in the active site titration experiment to obtain a near homogeneous ribozyme population. In addition, kinetic assays should be carried out by varying ribozyme/substrate ratio or by changing the concentration of substrate and ribozyme to ensure that burst population, burst rate, and product release rate remain constant under given reaction conditions. Finally, we also recommend using the assay to check the fraction of active population in each ribozyme stock to make sure that folding experiments are not conducted using degraded or nonactive RNAs. Kinetics as a tool to probe folding rate of a group II intron-derived ribozyme. The above multiple-turnover approach can be modified to determine the folding rate constant of a ribozyme construct. The reaction is set up such that folding, not chemistry, is rate-limiting during the first turnover [72]. The folding rate constant of D135 ribozyme was first determined from hydroxyl radical footprinting kinetics and found to be 1.0 min1 [73]. To validate this rate constant using multiple-turnover kinetics, we modified the setup of the assay to measure kfold as a function of catalysis. Here, substrate (10 lM final) and MgCl2 (100 mM final) are added concurrently to unfolded ribozyme (1 lM final) to start the reaction (Fig. 7; [73]). As such, observed rate from the first turnover is expected to reflect folding, not chemistry. Using this approach, we are able to observe a burst rate of 1 min1 , similar to the lower limit of kobs observed for the formation of active structure monitored via hydroxyl radical footprinting [73]. In addition, a burst amplitude of 100% is observed, suggesting that D135

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folded homogeneously into a single active conformation [73]. Assaying for the presence of a kinetic trap in the folding pathway. Given that D135 folds slowly to a single active conformation, an intriguing question is whether the ribozyme folds slowly due to the presence of a kinetic trap (as in most ribozymes [72,74–78]). The ability to monitor folding as a function of catalysis provides a useful way to address this question. To assay for kinetic traps, one determines how the folding rate is affected by the presence of denaturant [73]. If ribozyme folding is due to the presence of a kinetic trap, the addition of the denaturant generally accelerates folding rate constant by lowering the energetic barrier [75,78]. To examine this, we added excess substrate (10 lM final), MgCl2 (100 mM final), and subdenaturing concentration of urea (<1 M for D135 ribozyme) concurrently to the unfolded ribozyme (1 lM final) and assayed for product formation as described previously. The experiment was conducted at several different urea concentrations to determine how kobs changes with respect to [urea]. It was found that presence of urea does not accelerate the folding rate, suggesting that slow folding of D135 is not due to a kinetic trap [73].

6. Conclusion Kinetics is an essential biochemical tool for studying different mechanistic or structural aspects of RNA catalysis. Although kinetic assays are generally perceived as only a means for studying catalytic mechanisms, they also provide powerful tools for studying other aspects of ribozyme function, such as RNA folding, ribozyme specificity, fidelity, and conformational homogeneity of an RNA population.

Acknowledgments The authors thank Catherine Adamidi for helpful discussions on the manuscript. O.F. is a postdoctoral fellow at Howard Hughes Medical Institute. L.J.S acknowledges support from NIH Biophysics Training Grant (T32 GM08281). A.M.P is a Principal Investigator of the Howard Hughes Medical Institute.

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