CHEMICAL BIOLOGY

Group II Introns: Structure, Function, and Catalysis

 

Michael Roitzsch, Howard Hughes Medical Institute, Yale University, New Haven, Connecticut

doi: 10.1002/9780470048672.wecb679

 

The self-splicing group II introns are among the largest naturally occurring ribozymes and have been found primarily in organellar genomes of plants, fungi, bacteria, proteobacteria, and blue algae. More recently, a group II intron was found in a primitive metazoan. More than 200 different group II introns have been identified to date. All group II introns have a conserved secondary structure composed of six stem-loop structures termed domains D1 -D6, which are arranged around a central wheel and provide distinct contributions to intron function. Group II introns catalyze their own excision from pre-RNA by two consecutive transesterifications and concomitant splicing of the flanking exons. The reaction requires Mg2+ ions, which are directly involved in catalysis and are also needed for proper folding of the intron. Intriguingly, group II introns share several mechanistic and structural features with the eukaryotic spliceosome, which suggests an evolutionary relationship. In vivo, group II intron splicing is generally assisted by proteins. With the help of proteins, some group II introns can also reverse the splicing reaction and integrate themselves into target genomes;this process resembles transposition of non-LTR retrotransposons and can be exploited for biotechnological applications.

 

It was long believed that protein enzymes were the only cellular machines that could conduct biological catalysis. In 1982, however, Thomas R. Cech described the self splicing group I intron from the ciliated protozoon Tetrahymena thermophila, which can excise itself from pre-rRNA and join the flanking exons in vitro (1). Cech coined the term “ribozyme” for this new class of biocatalysts. At the same time, Sidney Altman reported that the RNA components of RNase P from Escherichia coli and Bacillus subtilis could cleave tRNA precursor molecules in the absence of protein cofactors (2). Over time, it has become obvious that ribozymes are widespread in nature. Ribozymes that catalyze a broad range of reactions have been detected in numerous different organisms, which include plants, lower eukaryotes, bacteria, and viruses (3). More recently, a ribozyme was even found in the human genome (4).

 

Background

Group II introns are a distinct subgroup within the naturally occurring ribozymes

With exception of the ribosome that catalyzes peptidyl transfer reactions, all naturally occurring ribozymes catalyze phosphoryl transesterifications on their parent RNA (in cis) or other RNAs (in trans) and can be roughly divided into three categories based on their catalytic properties.

1. Nucleolytic ribozymes. Many smaller (40-150 nucleotides) ribozymes catalyze site-specific cleavage in cis. Mechanistically, they activate a 2'-OH group to allow nucleophillic attack of the adjacent 3'-phosphate, which results in formation of a cyclic 2’,3’-phosphodiester and release of the downstream RNA strand (Fig. 1a). The reaction resembles nonspecific base-catalyzed RNA cleavage. Members of this category are the hammerhead ribozyme, the hepatitis delta virus ribozyme, the structurally related mammalian CPEB3 ribozyme, the hairpin ribozyme, the varkud satellite ribozyme, and the glmS riboswitch. Their function is to process replication intermediates or, in the case of the glmS riboswitch, the metabolite-dependent control of gene expression.

2. RNase P. RNase P is a trans-acting ribozyme (150-500 nucleotides) that processes pre-tRNAs by endonucleolytically cleaving off leader sequences from the 5’-end. RNase P positions and activates a water molecule to attack the scissile phosphate, which results in a free terminal 3’-OH group (Fig. 1b). Although they are usually associated with proteins, RNase P ribozymes have been shown to catalyze strand cleavage in the absence of their protein cofactors.

3. Self-splicing ribozymes. These ribozymes are noncoding intervening sequences (introns) that can catalyze their own excision from the parent pre-RNA. Two groups of self-splicing ribozymes exist: the group I and group II introns. They can be very large with sizes of 200-1500 nucleotides (group I introns) and 400-2500 nucleotides (group II introns). Ribozyme-catalyzed self-splicing is a two-step reaction.

In group I introns, the attacking nucleophile in the first step is the 3’-OH group of an external guanosine (Fig. 1c). In the second step of splicing, the 5’-exon terminus attacks the 3’-scissile phosphate, which results in spliced exons and a linear intron with a 5’-terminal guanosine.

In group II introns, the attacking nucleophile in the first step is either the 2’-OH group of an adenosine located close to the intron terminus (Fig. 1d) or a water molecule (analogous to Fig. 1b). The second step is similar to that of group I introns and involves attack of the 5’-exon terminus on the 3’-splice site and ligation of the two exons.

 

Figure 1. Schematic of phosphodiester bond-cleavage strategies used by different natural ribozymes. (a) In self-cleaving ribozymes, the 2’-OH group of the 3’-terminal nucleotide attacks the scissile phosphate, which results in a cyclic 2’,3’-phosphodiester. (b) RNase P activates a water molecule to allow a nucleophillic attack on the scissile phosphate, which forms a free 3’-OH group and a 5’-monophosphate at the resulting fragments. (c) In the first step of splicing of group I introns, the attacking nucleophile is provided by the 3’-OH group of a guanosine moiety. As a result, excised group I introns have an additional G at the 5’-terminus. (d) The attacking nucleophile in the first step of splicing of group II introns (branching pathway) is the 2’-OH group of a dedicated adenosine moiety (Abp) located close to the intron 3’-end in domain D6. This reaction results in a lariat form of the excised intron.

 

Similarities to the eukaryotic spliceosome

In higher organisms, splicing is catalyzed by a ribonucleoprotein complex termed spliceosome. Several mechanistic and structural similarities suggest that the spliceosome may have derived from ancestral group II introns. Both group II introns and the spliceosome catalyze splicing in a two-step reaction that results in formation of excised lariat introns, and they share the same stereospecific preferences at the splice sites (vide infra). In both systems, the two splicing steps proceed through inversion at the scissile phosphates, which is consistent with an SN2 reaction mechanism (5). More strikingly, the highly conserved group II intron domain D5 has metal ion-binding properties and structural features similar to the stem loop of the spliceosomal small nuclear RNA U6. These similarities are so substantial that an isolated domain (D5) from a group II intron can functionally replace a subsection (U6) of the spliceosomal RNA in an in vivo assay (6). Group II introns are therefore considered a valuable model for studying functions related to the more complex spliceosome.

Protein assisted splicing

Group II introns are essentially unreactive under physiological salt conditions (vide infra). Therefore, splicing in vivo requires protein cofactors (7). Various proteins have been associated with splicing. These proteins can be roughly divided into three groups.

1. Intron encoded maturases. These proteins, which are normally encoded by an open reading frame (ORF) within the intron, bind specifically their own RNA and promote formation of critical tertiary contacts. In addition, most of these proteins have endonuclease and reverse-transcriptase domains, which allow intron mobility (i.e., retrotransposition and retrohoming) by a mechanism termed target primed reverse transcription (TPRT) (vide infra).

2. Proteins recruited from the host. Many proteins have been associated with splicing, but most of these proteins cannot promote splicing in vitro, presumably because other cofactors are needed. However, two proteins, Cyt19 from Neurospora crassa and Mss116 from Saccharomyces cerevisiae, have recently been shown to promote splicing under near-physiological conditions in vitro. Both proteins belong to the family of DEAD-box helicases and promote splicing in an ATP dependent manner.

3. Proteins that promote splicing in an indirect way, for example by raising the intraorganellar Mg2+ concentration.

Reactions catalyzed by group II introns

Group II Intron-catalyzed self-splicing can proceed through two different pathways, which differ in the nucleophile that attacks in the first splicing step. In the branching pathway (Fig. 2a), this nucleophile is the 2'-OH group of the branch point adenosine, which is a bulged nucleotide located close to the intron terminus. This reaction leads to a covalently 2',5'-branched intron structure with a lariat form. The second reaction step involves attack of the 3'-terminal OH-group of the cleaved 5'-exon on the 3'-splice site. The reaction products are the excised lariat intron and the ligated exons.

The alternative splicing mode is the hydrolytic pathway (Fig. 2b). As the name indicates, a water molecule is the nucleophile that attacks the 5'-splice site in this reaction mode, which generates a linear intron. The second splicing step is identical to that of the branching pathway.

It is remarkable that group II introns can also bind the spliced exons and use them as substrate for different reactions. The simplest of these reactions is the spliced exon reopening, which is the irreversible hydrolytic cleavage of the exons at the original splice site (Fig. 2c).

A more interesting reaction is the complete reversal of both steps of splicing, which allow group II introns to reinsert themselves into the spliced exons (8). Reversal of the second step of splicing is an energetically neutral transesterification, which can be performed by linear and lariat introns. The situation is different for reversal of the first step of splicing, which has only been observed for the lariat intron. The lariat intron can reverse the first step of splicing by a second transesterification, which includes breaking of the 2',5'-phosphodiester at the branch point adenosine and formation of a new covalent linkage with the 5’-exon. The linear intron cannot perform a second transesterification because it lacks the 2’,5’-phosphodiester linkage. To join the 5’-exon to the intron, it would have to catalyze an energetically unfavorable condensation reaction, which has not been observed.

A striking feature of many group II introns is that they can also attack DNA substrates (9). With assistance of intron encoded maturases or other cofactors, they can insert themselves into double-stranded DNA by TPRT. In this reaction, the intron reverse splices into one DNA strand while the endonuclease domain of the maturase cuts the second strand. The maturase then uses the cleaved target DNA as a primer for reverse transcription, which generates a DNA copy of the intron. Group II introns can catalyze more reactions than those discussed here; for a more comprehensive overview on group II intron catalyzed reactions, see Reference 3.

 

Figure 2. (a) In the branching pathway the phosphate at the 5'-splice is attacked by the 2'-OH group of the branch point adenosine. In the second step of splicing, the 3'-terminal OH-group of the 5'-exon attacks the phosphate at the intron 3'-exon junction, which results in spliced exons and excised lariat intron. (b) In the hydrolytic pathway, the first step nucleophile is a water molecule that cleaves the 5'-intron, which forms a linear intron. The second step of splicing is essentially the same as in the branching pathway. (c) Spliced exon reopening requires binding of the substrate. The substrate is then irreversibly cleaved by activation of a water molecule.

 

Structure, Function, and Catalysis

Structural organization

Group II introns share a common secondary structure that is characterized by six stem-loop structures termed domains D1-D6 (3). The domains are arranged around a central wheel (Fig. 3). Group II introns can be divided into subgroups IIA, IIB, and IIC (10). The three subgroups share the consensus secondary structure but vary in the specific composition and lengths of individual domains. These variations reflect different ways to organize the overall tertiary structure of the intron, although the catalytic cores are presumably similar. In the catalytically active tertiary structure, the domains are closely packed together (7, 11). Several well-defined long-range interactions that connect the domains have been identified (Greek letters in Fig. 3), and most of them have been shown to be functionally essential.

 

 

Figure 3. All group II introns share a common secondary structure with six domains that radiate from a central wheel. Based on differences in some features, group II introns can be divided into subgroups IIA, IIB, and IIC. Long-range tertiary interactions are indicated by greek letters. The folding control element is indicated. The catalytic triad in domain D5 is marked by a rectangle. The branch point adenosine in domain D6 is shown as a bold ''A.'' Exons are depicted as gray boxes and intron binding sites are indicated (adapted from Reference 20).

 

Domain 1 (D1)

D1 is absolutely essential for catalytic activity. It provides a scaffold that helps the intron fold into the native tertiary structure. Several long-range interactions within D1, like the α-α', β-β', or the ω-ω' interaction, as well as contacts to other domains, which include ε-ε', K-K', λ-λ', θ-θ', and ς-ς', have been identified (Fig. 3). The λ-λ' and ε-ε' interactions are part of a larger, functional substructure that also involves the I(i) loop in the stem of D1. This substructure was named “z-anchor” and is important for intron function because it positions the 5'-splice site near the bulge of D5. In all group II introns, D1 contains the exon-binding site EBS1; this site is a short sequence that base pairs with a complementary sequence termed intron binding site (IBS) located at the 3'-end of the 5'-exon. Group IIA and IIB introns have an additional second stretch of nucleotides (EBS2) that interact with the 5' -exon. In addition, D1 contains a site termed EBS3 in group IIB and IIC and δ in group IIA introns, which binds to the first nucleotide of the 3'-exon. The EBS-IBS and, in group IIA introns, the δ-δ' interactions are of fundamental importance for the introns as they allow recognition and binding of the exons and determination of the splice site.

 

Domain 2 (D2)

In contrast to D1, D2 is not necessarily required for splicing, and most of this domain can be removed without severe effects on the catalytic activity. However, it stabilizes the tertiary structure by forming long-range interactions to D1 (θ-θ') and D6 (η-η’).

 

Domain 3 (D3)

D3 functions as a catalytic effector that strongly enhances the chemical rate of catalysis. To date, the μ-μ’ interaction is the only identified tertiary interaction between D3 and D5.

 

 

Domain 4 (D4)

D4 is not needed for splicing. In some introns, it contains an ORF encoding a maturase protein.

 

Domain 5 (D5)

D5 is a short stem-loop structure typically composed of 34 (group IIA and IIB) or 32 (group IIC) nucleotides with a conserved two-nucleotide bulge. It is the most conserved intronic domain and absolutely necessary for catalysis. It forms several contacts (λ-λ', K-K', and ς-ς') to D1 as well as a contact to D3 (μ-μ’). A very important feature of D5 is a highly conserved stretch of three consecutive nucleotides named the catalytic triad, which is located at the 5'-end of D5 and is absolutely required for catalysis. The sequence of the catalytic triad is AGC in group IIA and group IIB introns and CGC in group IIC introns. The catalytic triad and the nucleotides at the bulge form several direct contacts to divalent metal ions, which help to position them into the catalytic core.

 

Domain 6 (D6)

D6 is a hairpin structure that harbors the branch point adenosine, whose 2'-OH group acts as the nucleophile for the first step of splicing in the branching pathway. In addition, D6 contains an contact that forms the η-η' contact with D2. This element is a GNRA tetraloop-receptor interaction that is believed to mediate a conformational switch between the two steps of splicing.

 

J2/3 linker

Besides the six domains, the region joining domains D2 and D3 (J2/3) is particularly conserved and catalytically important. It forms the γ-γ' interaction, which joins J2/3 with the intron terminus and contains two consecutive nucleotides, GA in group IIA and IIB introns and GC in group IIC introns, which are critical for intron function. Photocrosslinking studies have shown that J2/3 is located in the proximity of the catalytic triad in D5. A recent crystal structure confirms these findings and shows that J2/3 forms base triplets with the catalytic triad and the bulge of D5, which helps to bring together catalytically important residues of the intron (12).

Trans splicing constructs and kinetic investigations

In vitro kinetic assays are an important tool to study the catalytic mechanisms of ribozymes. Intron mutant and deletion constructs have been used in kinetic assays to study the role of particular nucleotides and to distinguish whether a mutation causes a defect in chemistry or binding of exonic substrate or intronic components.

A very interesting feature of group II introns is their modularity; the intronic domains can be provided separately to form a functional ribozyme (9). For example, the exD123 construct consists of a 5'-exon and D1, D2, and D3. This construct alone is unreactive, but addition of the catalytically essential domain D5 in trans generates an active two-piece ribozyme that cleaves off the 5'-exon; it is therefore a mimic for the hydrolytic pathway of the first step of splicing. Similarly, an intron construct that consists of D5 and D6 (D56) can be combined with the exD123 construct to obtain a branching pathway mimic for the first step of splicing. To date, numerous other multipiece constructs have been successfully studied and some examples of naturally split group II introns have even been found plants (9). An important feature of trans splicing constructs in kinetic assays is that the individual constructs can be folded separately under appropriate conditions without reacting, and hence they allow the separation of folding from catalysis.

Group II introns are metal-dependent enzymes

Group II introns are metal-dependent enzymes that require Mg2+ for both folding and catalysis (9). Most group II introns are essentially unreactive at physiological salt concentrations. Hence, for in vitro experiments, nonphysiologically high salt concentrations, typically 500 mM of a monovalent and 100 mM Mg2+ are required to obtain optimal splicing conditions. In vivo, these high metal requirements are believed to be relieved by intron-encoded maturases and other cofactor proteins recruited from the host organism.

Metal ions have been localized by independent methods, which include phosphorothioate substitution (13), Tb3+ cleavage (14), X-ray crystallography (12), and NMR (15). Most identified metal contacts are provided by the phosphate backbone. Although the bridging phosphates are not chiral by chemical definition, the two nonbridging oxygen atoms are distinct in the tertiary structure. The nomenclature used to define these atoms is derived from phosphorothioate substitution experiments. Substitution of a single oxygen atom with sulfur at a bridging phosphate generates a chiral phosphorothioate; the two diastereomers are named as the Rp- and SP isomers according to the Cahn-Ingold-Prelog system. The nonbridging oxygens of a bridging phosphate can be identified accordingly as pro-SP and pro-RP oxygens.

 

Metal ions are required for folding

Studies with the group IIB intron ai5γ from the mitochondrial genome of S. cerevisiae have suggested that group II introns fold slowly into the native state via an on-pathway intermediate (16). In contrast, most other ribozymes proceed along a “rugged” folding pathway with stable misfolded intermediates (“kinetic traps”).

From the unfolded state, group II introns first fold slowly into a compact intermediate state and then, in a fast step, into the native state. At low Mg2+ concentrations, the introns can only reach the compact state, which was exploited to study this state in more detail. Using nucleotide analog interference mapping with compaction as the selection criterion, a small substructure in the heart of D1 was identified as a folding control element (Fig. 2). This element must adopt the correct conformation before other tertiary contacts of D1 can be established, leading to a highly compact tertiary structure of D1. Once the D1 scaffold is formed, high Mg2+ concentrations or a cofactor can facilitate rapid docking of the other domains into their designated binding sites thus forming the native (i.e., catalytically active) ribozyme.

 

Metal ions in the catalytic core

Many ribozymes and proteins that catalyze phosphoryl-transfer reactions use a mechanism employing two metal ions, and early on group II introns were hypothesized to use a similar mechanism (17). However, evidence for the existence of two metal ions in the catalytic core was only found very recently (12, 13, 18). A direct Mg2+ coordination of the pro-SP oxygen of the first nucleotide of the catalytic triad is evident based on phosphorothioate substitution experiments (18). Recently, an intact group II was successfully crystallized for the first time (12). This structure confirms the metal contact from the first nucleotide of the catalytic triad and shows additional contacts to this metal ion from the second nucleotide of the catalytic triad and from the first nucleotide upstream of the bulge (Fig. 4a). This nucleotide is additionally coordinated to the second Mg2+ ion in the core. The distance between the two Mg2+ ions in the crystal structure is 3.9 A, which is in agreement with the proposed two-metal-ion mechanism (17).

Because the data from phosphorothioate substitution experiments, which provide information about the metal binding situation before splicing, match the crystal structure that captures the situation after splicing, it may be concluded that the two metal ions are tightly bound in the catalytic center in an arrangement that remains essentially unchanged through both steps of splicing.

During splicing, the two metal ions form additional contacts to the substrate. Evident from phosphorothioate substitution experiments are coordinative bonds to the pro-RP oxygens of the scissile phosphates at both splice sites and to the 3'-oxygens of the leaving groups in both steps of splicing (Fig. 4).

 

 

Figure 4. Chemical mechanism of group II intron splicing (branching pathway). The sequences and numbering are according to the ai5γ intron from S. cerevisiae. The solid arrows between the panels mark the forward splicing direction, the dotted arrows the reverse splicing direction. (a) The first step is initiated by attack of the 2-OH group of the branch point adenosine (A880) at the scissile phosphate as indicated by the arrow. The dotted lines indicate coordinative bonds to metal ions. Metal ion coordinations from the catalytic triad (A816-C818) and the bulge in D5 (C837-C839) are indicated in this panel but omitted in the following panels for clarity. (b) In the transition state of the first step of splicing, the scissile phosphate is expected to have a trigonal bipyramidal arrangement. (c) In the splicing intermediate, the metal coordinated oxygen of the 5’-scissile phosphate has formally changed from pro-Rp to pro-Sp. The second step of splicing is initiated by attack of the 3’-terminal oxygen of the 5’-exon at the 3’-scissile phosphate as indicated by the solid arrow. The dotted arrow indicates the reverse splicing reaction. (d) The transition state of the second step of splicing is expected to have a trigonal bipyramidal arrangement at the scissile phosphate similar to the transition state of the first step of splicing. (e) Schematic of the products immediately after the second chemical step. This arrangement is also the initial configuration expected for the first step of reverse splicing as indicated by the dotted arrow.

 

The chemical mechanism of group II intron splicing

The following paragraph will discuss the detailed chemical mechanism of both steps of splicing by the branching pathway. Before the first chemical step of splicing takes place, the substrate is properly arranged in the catalytic core, which is mainly facilitated by the EBS-IBS and δ-δ' interactions, and the metal ions are positioned as described above (Fig. 4a).

 

The first step of splicing

In the first step of splicing, the 2'-OH group of the branch point adenosine attacks the scissile phosphate at the designated 5'-splice site. It is not established how the nucleophile is activated; however, it is likely that the 2'-OH is coordinated to a metal ion in the core that could facilitate deprotonation (17).

Both steps of splicing are known to proceed with inversion at the scissile phosphates, which is consistent with an SN2 substitution (5). In the transition state, the scissile phosphate is therefore expected to adopt a trigonal bipyramidal arrangement (Fig. 4b). Metal ion coordination to a nonbridging oxygen of this phosphate might help to compensate the emerging negative charge in the transition state. The leaving group (i.e., the 3’-oxygen of the 5’-exon) is likewise stabilized by metal ion coordination. In the resulting intermediate, the metal-ion coordinated oxygen atom of the 5'-splice site phosphate has formally changed from pro-RP to pro-SP because of the inversion (Fig. 4c).

 

The second step of splicing

The leaving group of the first step of splicing, which is the 3'-terminal OH group of the 5'-exon, becomes the attacking nucleophile in the second step of splicing. Metal-coordination properties of the participating reaction centers are comparable to the first step. The pro-RP oxygen of the scissile phosphate, the 3’-oxygen of the 3’-terminal intron nucleotide (the leaving group) and the 3'-terminal nucleotide of the 5’-exon (the attacking nucleophile) are coordinated to divalent metal ions (Fig. 4c).

The transition state of the second step of splicing resembles that of the first step. The scissile phosphate adopts a trigonal bipyramidal arrangement and is stabilized by metal coordination to a non-bridging oxygen atom (Fig. 4d). The leaving group, i.e. the 3'-oxygen of the terminal intron nucleotide, is likewise stabilized by metal coordination. The resulting products are the lariat intron and the spliced exons (Fig. 4e). This arrangement is also the starting conformation for the reverse splicing reaction, which is indicated by dotted arrows in Fig. 4.

Future Research Topics

The story of group II introns is not at the end. The recent crystal structure of an intact group II intron will presumably lead to several follow-up studies. For example, it would be interesting to crystallize and compare group II introns from different subgroups or to capture different reaction states of the intron.

Group II introns naturally catalyze a broad range of reactions. The catalytic repertoire of other ribozymes was previously extended by in vitro evolution. It could be very exciting to apply this method to the versatile group II introns.

The ability to invade duplex DNA has led to useful biotechnological applications. By manipulating the exon-binding sites in domain D1 of the intron, which recognize potential target sites, group II introns can be reprogrammed to attack a given sequence selectively. This method was used to develop group II introns into gene-targeting vectors (“targetrons”) (19). Targetrons may have a great potential for medical applications. They could be used to knock out or to repair a malfunctioning gene of choice. Another idea is to engineer the intron ORF in D4 to contain additional coding sequences; group II introns could then be used to introduce new genes into an organism.

Acknowledgments

The author thanks Anna M. Pyle, Amanda Solem, Nora Zingler, and Olga Fedorova for critical reading of the manuscript and useful discussions.

References

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6. Shukla GC, Padgett RA. A catalytically active group II intron domain 5 can function in the U12-dependent spliceosome. Mol. Cell 2002; 9:1145-1150.

7. Solem A, Zingler N, Pyle AM, Li-Pook-Than J. Group II introns and their protein collaborators. In: Non-Protein Coding RNAs. Walter NG, Woodson SA, Batey RT, eds. 2008. Springer, Berlin.

8. Augustin S, Muller MW, Schweyen RJ. Reverse self-splicing of group II intron RNAs in vitro. Nature 1990; 343:383-386.

9. Pyle AM. Group II introns: catalysts for splicing, genomic change and evolution. In: Ribozymes and RNA Catalysis. Lilley DMJ, Eckstein F, eds. 2008. RCS Publishing, Cambridge, UK.

10. Toor N, Hausner G, Zimmerly S. Coevolution of group II intron RNA structures with their intron-encoded reverse transcriptases. RNA 2001; 7:1142-1152.

11. Swisher J, Duarte CM, Su LJ, Pyle AM. Visualizing the solvent-inaccessible core of a group II intron ribozyme. EMBO J. 2001; 20:2051-2061.

12. Toor N, Keating KS, Taylor SD, Pyle AM. Crystal structure of a self-spliced group II intron. Science 2008; 320:77-82.

13. Gordon PM, Fong R, Piccirilli JA. A second divalent metal ion in the group II intron reaction center. Chem. Biol. 2007; 14:607-612.

14. Sigel RK, Vaidya A, Pyle AM. Metal ion binding sites in a group II intron core. Nat. Struct. Biol. 2000; 7:1111-1116.

15. Erat MC, Sigel RK. Determination of the intrinsic affinities of multiple site-specific Mg(2+) ions coordinated to domain 6 of a group II intron ribozyme. Inorg. Chem. 2007; 46:11224-11234.

16. Pyle AM, Fedorova O, Waldsich C. Folding of group II introns: a model system for large, multidomain RNAs? Trends Biochem. Sci. 2007; 32:138-145.

17. Steitz TA, Steitz JA. A general two-metal-ion mechanism for catalytic RNA. Proc. Natl. Acad. Sci. U.S.A. 1993; 90:6498-6502.

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Further Reading

Gesteland RF, Cech TR, Atkins JF, eds. The RNA World, 3rd ed. 2006.

Cold Spring Harbor Lab. Press, Cold Spring Harbor, NY.

Lambowitz AM, Zimmerly S. Mobile Group II Introns. Annu. Rev. Genet. 2004; 38:1-35.

Lilley DMJ, Eckstein F, eds. Ribozymes and RNA Catalysis. 2008. RCS Publishing, Cambridge, UK. http://www.fp.ucalgary.ca/group2introns/.

 

See Also

Bacterial DNA polymerases, chemistry of

Origins of life: emergence of the RNA world

RNA, noncoding