Catalytic RNA


Ribonucleic acid (RNA) molecules have diverse roles in biological systems. Although some code for proteins or act to translate codons to amino acids, others fold into specific shapes that endow them with the ability to catalyse specific chemical transformations. These catalytic RNAs, ribozymes, are responsible for protein synthesis, transfer RNA (tRNA) processing, self‐splicing of certain introns, self‐scission during rolling circle replication of some single‐stranded RNA viruses and cofactor‐dependent gene regulation in bacteria. Other ribozymes have been evolved in vitro to perform a wide variety of transformations. Two of these, tRNA aminoacylase and RNA polymerase ribozymes, are featured here because molecules with such capabilities are thought to have existed on early Earth, before proteins took over as the dominant biological catalysts. Most of the ribozymes have been shown to perform multiturnover catalysis and thus act as true enzymes, either in their natural, biological form or as engineered constructs.

Key Concepts:

  • RNA molecules can fold into specific conformations and accelerate chemical transformations, thus acting as catalytic biomacromolecules, ribozymes.

  • Ribozymes can accelerate chemical reactions by many orders of magnitude.

  • Many ribozymes are capable of multiturnover catalysis, acting as enzymes.

  • Protein synthesis templated by mRNA is catalysed by ribosomal RNA.

  • Most ribozymes are phosphoryl transferases.

  • In vitro selected ribozymes have been shown to catalyse a wide variety of reactions.

  • The existence of ribozymes supports the RNA World hypothesis.

Keywords: ribozyme; RNA secondary structure; in vitro selection; acid–base mechanism; polymerisation; metal ion

Figure 1.

Structures and proposed transition state of the ribosome (a–c) and RNase P (d–g). (a) The ribosome is responsible for mRNA‐templated protein synthesis in all cellular life forms and is composed of both RNA (dark blue) and protein domains (light blue). (b) For accurate and efficient protein synthesis, the ribosome binds mRNA (dark grey) and translates the codons into amino acids (decoding site). The ribosome then binds the correct tRNA (blue) corresponding to the codons and preforms a peptidyl transferase reaction to increase the length of the nascent peptide (red) by one amino acid. (c) The carbonyl carbon that links the nascent peptide to the tRNA at the P‐site undergoes a nucleophilic attack by the A‐site‐bound aminoactyl‐tRNA. The proposed transition state was determined by kinetic isotope‐effect analysis and shows a two step mechanism in which the proton donation from the nucleophilic nitrogen and the formation of the tetrahedral intermediate are both in the rate limiting step with the fast break down of the tetrahedral intermediate following. (d) Crystal structure of the Thermotoga maritima RNase P holoenzyme (S‐ (green) and C‐ (blue) domains and a small protein cofactor (tan)) in complex with processed tRNA (red). The ribozyme‐product complex illustrates the intermolecular contacts necessary for substrate binding and processing. (e) The secondary structure of RNase P denotes the S‐ and C‐domains (shades of green and blue, respectively) along with the tRNA identification sites (red) and contacts (grey boxes). The 5′ end of the tRNA marks the location of the active site near stems P4 and P5, whereas the 3′ terminus is base‐paired with L15 and is necessary for identification of the particular type of tRNA. (f) RNase P is mainly responsible for the processing of the 5′ ends of tRNA, although it does process other RNAs. A hydroxide, or metal‐coordinated hydroxide, acts as the nucleophile attacking the phosphate at the cleavage site, resulting in a 3′‐hydroxyl on the 5′‐leader and a 5′‐phosphate on the processed tRNA. (g) Two metal ions have been proposed to stabilise the transition state. The first metal (M1) is proposed to interact with the nonbridging scissile phosphate oxygen allowing the nucleophilic hydroxide to attack. The second metal (M2) is necessary for stabilisation of the transition state and proton donation to the 3′‐hydroxyl on the 5′ product (Reiter et al., ).

Figure 2.

Structures and proposed transition state of the group I (a–d) and group II (e–h) self‐splicing introns. (a) Crystal structure of the purple bacterium, Azoarcus group I, self‐splicing intron in complex with both the 5′‐ and 3′‐ exons from the tRNA anticodon loop. This structure corresponds to the intermediate after the first of two consecutive transesterification reactions. The crystal structure shows the 3′ hydroxyl of the 5′ exon in line for nucleophilic attack on the scissile phosphate, which results in the excision of the intron and ligation of the exons. The catalytic residues are explicitly shown in the active site. (b) There are three coaxially stacked helical elements: P5, P4, P6 and P6a (green); P10, P1 and P2 (orange); P9.0, P7, P3 and P8 (blue). The 3′ and 5′ exons are depicted in red and yellow boxes with lower‐case letters and are base‐paired with P10 and P1, respectively (Adams et al., ). The splice site is shown as a grey arrowhead in both the secondary and crystal structures. (c) The splicing reaction proceeds through two transesterification reactions in which an exogenous guanosine attacks the 5′ splice site and generates a new 3′–5′ phosphodiester bond. The resulting 5′ exon contains a free 3′ hydroxyl that is available for attack on the 3′ splice site, completing the splicing reaction. (d) The proposed transition state for the second transesterification reaction, corresponding to the crystal and secondary structures, is shown with two metal ions coordinated in the active site yielding the ligated exons (Shan et al., ; Stahley and Strobel, ). (e) Crystal structure of the group II intron from Oceanobacillus iheyensis in the pre‐catalytic state. The G359A mutant abolishes catalytic activity by changing the conserved G·U wobble to an A–U base pair, while maintaining similar overall structure to the post‐catalytic state and represents the first crystal structure of an intron before splicing. The catalytic triad and A376 are explicitly shown, as well as the 5′ exon (tan), and the cleavage site with a grey arrowhead (Chan et al., ). (f) The ribozyme contains six domains (DI–DVI) with DI (green) being the largest. The most highly conserved structure and catalytic core are contained within DV (red). This includes a hairpin secondary structure and the ‘catalytic triad’ of conserved nucleotides and is located in close proximity to the catalytic bulged adenosine in DVI that acts as the nucleophile (DVI is not shown in the crystal structure). (g) Two transesterification reactions are necessary for group II intron splicing. The first is initiated by a DVI bulged adenosine that acts as a nucleophile in the 5′ splice reaction using its 2′ hydroxyl to liberate a 3′ hydroxyl which then attacks the 3′ splice site, resulting in ligation of the exons and excision of the intron. (h) The proposed transition state for the second transesterification reaction in which at least two metal ions are coordinated in the active site.

Figure 3.

Structures and proposed transition state of the hepatitis delta virus and hammerhead ribozymes. (a) Crystal structure of the HDV ribozyme in the pre‐cleaved state. To inhibit the ribozyme self‐cleavage and to capture the pre‐cleavage structure, the ribozyme is bound to an inhibitor RNA that contains a deoxynucleotide at the cleavage site (Chen et al., ). (b) This ribozyme folds into the complex nested‐double pseudoknot structure with two coaxial helices; P1 stacked with P1.1 and P4 (shades of blue) and P2 stacked with P3 (shades of red) depicted in the secondary structure. An active site cytosine is shown explicitly as is a divalent metal ion with the site of self‐scission indicated by a grey arrowhead. (c) The self‐cleavage reaction most likely proceeds through a general acid–base mechanism. A hydrated magnesium ion is suggested to act as a general base, deprotonating the 2′ hydroxyl upstream of the cleavage site and stabilising the phosphorene transition state. This coordination allows for nucleophilic attack on the phosphate backbone, generating a 2′–3′ cyclic phosphate on the 5′ product and a 5′ hydroxyl on the 3′ product. The C75 nucleobase likely acts as the general acid, protonating the leaving group. (d) Crystal structure of the full‐length Schistosoma mansonihammerhead ribozyme. The full‐length ribozyme reveals tertiary contacts between stems I and II that account for the 1000‐fold catalytic enhancement relative to the minimal sequence (Martick and Scott, ). (e) A general secondary structure denoting the fold of the hammerhead ribozyme. There are three classes – Type I, II and III – depending on which stem is open to the remainder of the transcript. Catalytic residues G12 and G8 are shaded in grey in the secondary structure and explicitly shown in the crystal structure along with a grey arrowhead indicating the site of cleavage in both. (f) The self‐scission proceeds through a general acid–base pathway in which the conserved G12 residue has direct contact with the cleavage site and acts as the general base deprotonating the 2′ hydroxyl. The conserved G8 residue is hydrogen bonded to the scissile phosphate, providing stabilisation of the transition state. G8 may also act as a general acid donating a proton to the leaving group.

Figure 4.

Structures and proposed transition state of the hairpin and glmS ribozymes. (a) Crystal structure of the hairpin ribozyme in the pre‐cleavage form. The ribozyme is trapped in the precursor state by substitution of the nucleophilic 2′ hydroxyl with a methoxy group, retaining structure but abolishing self‐cleavage activity. (b) Secondary structure of the hairpin ribozyme. The ribozyme consists of four helical stems centred on a four‐way junction. There are two coaxial stacks with stem A stacked on stem D (shades of red) and stem C stacked on stem B (shades of blue). Stems A and B are docked. (c) The active form of the ribozyme catalyses reversible transesterification reactions of site‐specific self‐cleavage and ligation. The proposed transition state for the self‐cleavage reaction is shown with a vanadate transition state analogue. Nucleobase G8 in loop A hydrogen bonds with the nucleophile, suggesting that it acts as the general base deprotonating the 2′‐hydroxyl initiating the reaction. The transition state is proposed to be stabilised by nucleobase A38 through hydrogen bonding and it is also suggested to act as the general acid donating a proton to the leaving group (Rupert et al., ). (d) Crystal structure of the Bacillus anthracis glmS ribozyme in complex with substrate gluclosamine‐6‐phosphate (GlcN6P). The structure contains an active site G40A mutation abolishing self‐cleavage activity but retaining overall structure. GlcN6P is explicitly shown in the crystal structure along with catalytic residue G40A. (e) The secondary structure of the glmS ribozyme consists of three coaxially stacked helices with P1 stacked on P2.2 (shades of blue), P2 stacked on P3 (shades of red) and P2.1 stacked on P3.1 (shades of green) arranged in a double pseudoknot. The catalytic core is located in P2.1 and P2.2 with six nucleotides responsible for binding the cofactor GlcN6P, outlined in black, and the catalytic residue G40, shaded in grey. The cleavage site is indicated with a grey arrowhead. (f) The deprotonated N1 on nucleobase G40 is predicted to act as the general base, deprotonating the 2′‐hydroxyl at the cleavage site. The general acid is suggested to be the protonated form of GlcN6P, which donates a proton to the leaving group, completing the self‐cleavage reaction (Klein and Ferre‐D'Amare, ).

Figure 5.

Structures and proposed reaction mechanism of the in vitro selected flexizyme and RNA polymerase ribozymes. (a) Crystal structure of the flexizyme, which preforms ribozyme‐catalysed tRNA aminoacylation, fused to an RNA substrate. Phenylalanine and a magnesium ion are shown in the binding pocket. (b) The core consists of two internal helices, P1 and P2, with an additional three base‐pair recognition helix formed between the 3′ end of the flexizyme and the tRNA in P3. (c) The aminoacylation occurs in two steps. First, the 5′ hydroxyl of the G1 position of the ribozyme self‐aminoacylates utilising an activated phenylalanine derivative. Second, the charged ribozyme recognises the correct tRNA substrate via base‐pairing and exclusively aminoacylates its terminal 3′ hydroxyl completing the reaction (Xiao et al., ). (d) Crystal structure of the post‐ligation state of the ligase ribozyme that functions as the core of the polymerase ribozyme (Shechner et al., ). (e) The core of the in vitro selected RNA‐dependent RNA ligase contains three coaxially stacked domains with P2 stacked on P1 (shades of green), P3 stacked on P6 and P7 (shades of blue) and P4 stacked on P5 (shades of red). The ligation site is contained within P1, whereas P4 and the catalytic C47, shown shaded in grey, for the active site. The site of ligation is indicated by a grey arrowhead. (f) The ligation reaction is initiated by an attack of the α‐phosphate at the 5′ end of the ribozyme by the 3′ hydroxyl on the substrate, ligating the substrate and releasing pyrophosphate. The suggested transition state is shown with the ligation substrate in blue. One magnesium ion is shown in the active site, possibly stabilising the transition state (Shechner and Bartel, ).



Adams PL, Stahley MR, Kosek AB, Wang J and Strobel SA (2004) Crystal structure of a self‐splicing group I intron with both exons. Nature 430: 45–50.

Bartel DP and Szostak JW (1993) Isolation of new ribozymes from a large pool of random sequences. Science 261: 1411–1418.

Chan RT, Robart AR, Rajashankar KR, Pyle AM and Toor N (2012) Crystal structure of a group II intron in the pre‐catalytic state. Nature Structural and Molecular Biology 19: 555–557.

Chen JH, Yajima R, Chadalavada DM et al. (2010) A 1.9 A crystal structure of the HDV ribozyme precleavage suggests both Lewis acid and general acid mechanisms contribute to phosphodiester cleavage. Biochemistry 49: 6508–6518.

Cochrane JC, Lipchock SV and Strobel SA (2007) Structural investigation of the GlmS ribozyme bound to its catalytic cofactor. Chemistry and Biology 14: 97–105.

De la Pena M, Gago S and Flores R (2003) Peripheral regions of natural hammerhead ribozymes greatly increase their self‐cleavage activity. EMBO Journal 22: 5561–5570.

Ekland EH, Szostak JW and Bartel DP (1995) Structurally complex and highly active RNA ligases derived from random RNA sequences. Science 269: 364–370.

Ferre‐D'Amare AR and Scott WG (2010) Small self‐cleaving ribozymes. Cold Spring Harbor Perspectives in Biology 2: a003574. doi:10.1101/cshperspect.a003574.

Ferre‐D'Amare AR, Zhou K and Doudna JA (1998) Crystal structure of a hepatitis delta virus ribozyme. Nature 395: 567–574.

Golden BL (2011) Two distinct catalytic strategies in the hepatitis delta virus ribozyme cleavage reaction. Biochemistry 50: 9424–9433.

Guerrier‐Takada C, Gardiner K, Marsh T, Pace N and Altman S (1983) The RNA moiety of ribonuclease‐P is the catalytic subunit of the enzyme. Cell 35: 849–857.

Guo HC and Collins RA (1995) Efficient trans‐cleavage of a stem‐loop RNA substrate by a ribozyme derived from Neurospora VS RNA. EMBO Journal 14: 368–376.

Hammann C, Luptak A, Perreault J and de la Pena M (2012) The ubiquitous hammerhead ribozyme. RNA – A Publication of the RNA Society 18: 871–885.

Hiller DA, Singh V, Zhong M and Strobel SA (2011) A two‐step chemical mechanism for ribosome‐catalysed peptide bond formation. Nature 476: 236–239.

Johnston WK, Unrau PJ, Lawrence MS, Glasner ME and Bartel DP (2001) RNA‐catalyzed RNA polymerization: accurate and general RNA‐templated primer extension. Science 292: 1319–1325.

Khvorova A, Lescoute A, Westhof E and Jayasena SD (2003) Sequence elements outside the hammerhead ribozyme catalytic core enable intracellular activity. Nature Structural Biology 10: 708–712.

Klein DJ and Ferre‐D'Amare AR (2006) Structural basis of glmS ribozyme activation by glucosamine‐6‐phosphate. Science 313: 1752–1756.

Kruger K, Grabowski PJ, Zaug AJ et al. (1982) Self‐splicing RNA – Auto‐excision and auto‐cyclization of the ribosomal‐RNA intervening sequence of tetrahymena. Cell 31: 147–157.

Kuhlenkoetter S, Wintermeyer W and Rodnina MV (2011) Different substrate‐dependent transition states in the active site of the ribosome. Nature 476: 351–354.

Lambowitz AM and Zimmerly S (2011) Group II introns: mobile ribozymes that invade DNA. Cold Spring Harbor Perspectives in Biology 3: a003616.

Leclerc F (2010) Hammerhead ribozymes: True metal or nucleobase catalysis? Where is the catalytic power from? Molecules 15: 5389–5407.

Lee N, Bessho Y, Wei K, Szostak JW and Suga H (2000) Ribozyme‐catalyzed tRNA aminoacylation. Nature Structural Biology 7: 28–33.

Lipfert J, Ouellet J, Norman DG, Doniach S and Lilley DM (2008) The complete VS ribozyme in solution studied by small‐angle X‐ray scattering. Structure 16: 1357–1367.

Martick M and Scott WG (2006) Tertiary contacts distant from the active site prime a ribozyme for catalysis. Cell 126: 309–320.

Mcclain WH, Guerriertakada C and Altman S (1987) Model substrates for an RNA enzyme. Science 238: 527–530.

Nissen P, Hansen J, Ban N, Moore PB and Steitz TA (2000) The structural basis of ribosome activity in peptide bond synthesis. Science 289: 920–930.

Noller HF, Hoffarth V and Zimniak L (1992) Unusual resistance of peptidyl transferase to protein extraction procedures. Science 256: 1416–1419.

Peebles CL, Perlman PS, Mecklenburg KL et al. (1986) A self‐splicing RNA excises an intron lariat. Cell 44: 213–223.

Reiner R, Ben‐Asouli Y, Krilovetzky I and Jarrous N (2006) A role for the catalytic ribonucleoprotein RNase P in RNA polymerase III transcription. Genes and Development 20: 1621–1635.

Reiter NJ, Osterman A, Torres‐Larios A et al. (2010) Structure of a bacterial ribonuclease P holoenzyme in complex with tRNA. Nature 468: 784–789.

Rupert PB and Ferre‐D'Amare AR (2001) Crystal structure of a hairpin ribozyme‐inhibitor complex with implications for catalysis. Nature 410: 780–786.

Rupert PB, Massey AP, Sigurdsson ST and Ferre‐D'Amare AR (2002) Transition state stabilization by a catalytic RNA. Science 298: 1421–1424.

Salehi‐Ashtiani K, Luptak A, Litovchick A and Szostak JW (2006) A genomewide search for ribozymes reveals an HDV‐like sequence in the human CPEB3 gene. Science 313: 1788–1792.

Schmelzer C and Schweyen RJ (1986) Self‐splicing of group II introns in vitro: mapping of the branch point and mutational inhibition of lariat formation. Cell 46: 557–565.

Schon A (1999) Ribonuclease P: the diversity of a ubiquitous RNA processing enzyme. FEMS Microbiology Reviews 23: 391–406.

Shan S, Kravchuk AV, Piccirilli JA and Herschlag D (2001) Defining the catalytic metal ion interactions in the tetrahymena ribozyme reaction. Biochemistry 40: 5161–5171.

Shechner DM and Bartel DP (2011) The structural basis of RNA‐catalyzed RNA polymerization. Nature Structural and Molecular Biology 18: 1036–1042.

Shechner DM, Grant RA, Bagby SC et al. (2009) Crystal structure of the catalytic core of an RNA‐polymerase ribozyme. Science 326: 1271–1275.

Sievers A, Beringer M, Rodnina MV and Wolfenden R (2004) The ribosome as an entropy trap. Proceedings of the National Academy of Sciences of the USA 101: 7897–7901.

Stahley MR and Strobel SA (2005) Structural evidence for a two‐metal‐ion mechanism of group I intron splicing. Science 309: 1587–1590.

Toor N, Keating KS, Taylor SD and Pyle AM (2008) Crystal structure of a self‐spliced group II intron. Science 320: 77–82.

Valadkhan S, Mohammadi A, Jaladat Y and Geisler S (2009) Protein‐free small nuclear RNAs catalyze a two‐step splicing reaction. Proceedings of the National Academy of Sciences of the USA 106: 11901–11906.

van der Veen R, Arnberg AC, van der Horst G et al. (1986) Excised group II introns in yeast mitochondria are lariats and can be formed by self‐splicing in vitro. Cell 44: 225–234.

Veeraraghavan N, Ganguly A, Chen JH et al. (2011) Metal binding motif in the active site of the HDV ribozyme binds divalent and monovalent ions. Biochemistry 50: 2672–2682.

Webb CH, Riccitelli NJ, Ruminski DJ and Luptak A (2009) Widespread occurrence of self‐cleaving ribozymes. Science 326: 953.

Wilson DS and Szostak JW (1999) In vitro selection of functional nucleic acids. Annual Review of Biochemistry 68: 611–647.

Winkler WC, Nahvi A, Roth A, Collins JA and Breaker RR (2004) Control of gene expression by a natural metabolite‐responsive ribozyme. Nature 428: 281–286.

Wochner A, Attwater J, Coulson A and Holliger P (2011) Ribozyme‐catalyzed transcription of an active ribozyme. Science 332: 209–212.

Xiao H, Murakami H, Suga H and Ferre‐D'Amare AR (2008) Structural basis of specific tRNA aminoacylation by a small in vitro selected ribozyme. Nature 454: 358–361.

Zaug AJ, Grabowski PJ and Cech TR (1983) Autocatalytic cyclization of an excised intervening sequence RNA is a cleavage‐ligation reaction. Nature 301: 578–583.

Further Reading

Das SR and Piccirilli JA (2005) General acid catalysis by the hepatitis delta virus ribozyme. Nature Chemical Biology 1: 45–52.

Fedor MJ and Williamson JR (2005) The catalytic diversity of RNAs. Nature Reviews Molecular Cell Biology 6: 399–412.

Gesteland RF and Atkins JF (eds) (1993) The RNA world. In: Cold Spring Harbor, Monograph Series, vol. 24. New York: Cold Spring Harbor Press.

Leung EK, Suslov N, Tuttle N, Sengupta R and Piccirilli JA (2011) The mechanism of peptidyl transfer catalysis by the ribosome. Annual Review of Biochemistry 80: 527–555.

Lonnberg T (2011) Understanding catalysis of phosphate‐transfer reactions by the large ribozymes. Chemistry – A European Journal 17: 7140–7153.

Ramakrishnan V (2002) Ribosome structure and the mechanism of translation. Cell 108: 557–572.

Webb CH and Luptak A (2011) HDV‐like self‐cleaving ribozymes. RNA Biology 8: 719–727.

Wilson TJ, Nahas M, Ha T and Lilley DM (2005) Folding and catalysis of the hairpin ribozyme. Biochemical Society Transactions 33: 461–465.

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Burke, Cassandra R, and Lupták, Andrej(Dec 2012) Catalytic RNA. In: eLS. John Wiley & Sons Ltd, Chichester. [doi: 10.1002/9780470015902.a0000870.pub2]