TY - JOUR
AU - Berzal-Herranz,, Alfredo
AB - Abstract The discovery 20 years ago that some RNA molecules, called ribozymes, are able to catalyze chemical reactions was a breakthrough in biology. Over the last two decades numerous natural RNA motifs endowed with catalytic activity have been described. They all fit within a few well-defined types that respond to a specific RNA structure. The prototype catalytic domain of each one has been engineered to generate trans-acting ribozymes that catalyze the site-specific cleavage of other RNA molecules. On the 20th anniversary of ribozyme discovery we briefly summarize the main features of the different natural catalytic RNAs. We also describe progress towards developing strategies to ensure an efficient ribozyme-based technology, dedicating special attention to the ones aimed to achieve a new generation of therapeutic agents. Ribozyme, Catalytic RNA, RNA-based tool, Gene silencing 1 Introduction Ribozymes are RNA molecules with catalytic activity. Since their discovery in the early eighties [1], RNA chemical catalysis has been described in several essential biological processes such as RNA splicing, RNA processing, the replication of RNA genomes, and peptide bond formation during translation [2–5]. Naturally occurring ribozymes, with the sole exception of the ribosome, catalyze the cleavage or ligation of the RNA phosphodiester backbone. In contrast to other known ribonucleases, ribozymes catalyze highly sequence-specific reactions determined by RNA–RNA interactions between the ribozyme and its substrate molecules. The key to the recognition of and binding to the substrate molecule, and the cleavage reaction, resides in the RNA molecule. The capacity of ribozymes to specifically inactivate other RNAs has made ribozymes very promising molecular tools and potential gene suppressors with important applications. Most of the naturally occurring ribozymes (except Ribonuclease P (RNase P) and ribosomes) catalyze intramolecular reactions in their natural context [1, 6–8]. The catalytic center of these naturally self-cleaving ribozymes has been identified and engineered to develop agents that catalyze intermolecular reactions, i.e., ribozymes that cleave external RNA molecules [7, 9–11]. These behave like true enzymes: they are not modified during the reaction and one ribozyme molecule can process several substrate molecules. Substrate recognition and binding is essentially governed by Watson–Crick interactions. Since, in general, the ribozyme sequences involved in substrate recognition are not involved in the chemical step of the reaction, the specificity of ribozymes can easily be changed to target virtually any RNA of interest. Numerous studies have tried to test the ability of ribozymes to inactivate cellular or viral RNAs, and to develop strategies for using ribozymes within the cell milieu. These studies have proven the usefulness of ribozymes as specific gene silencing tools, and have shown their potential in the development of new therapeutic agents, etc. This article briefly reviews the main features of different natural ribozymes, and looks at the work undertaken to develop RNA tools from their catalytic domains. It also discusses the many reports describing the use of ribozymes as specific suppressors of gene function, and the different strategies used to improve ribozyme catalytic performance in mammalian cells. Finally, the potential application of these molecules, based on such studies, is examined. 2 General features In this section we will go through the general features of the different naturally occurring ribozymes (with the exception of the ribosome). They can be separated into three groups: self splicing introns, RNase P and small catalytic RNAs. Two classes of self-splicing introns can be distinguished (group I and group II) based on their splicing mechanism and secondary structure. RNase P is an ubiquitous enzyme involved in the biosynthesis of tRNAs. Finally, the small catalytic RNA group includes four different catalytic motifs that share a common chemical reaction. In this group we can distinguish the so called hammerhead, hairpin, hepatitis delta virus (HDV) and VS ribozymes. 2.1 Self-splicing introns RNA splicing is an essential process in the expression of many genes. It consists of the precise excision of the intron and the covalent linkage of the boundary exons. A large number of introns belonging to group I or group II have been shown to catalyze their own splicing in vitro [12]. 2.1.1 Group I introns The first hint of catalytic activity in RNA molecules concerned a group I intron found in the precursor of Tetrahymena termophila large subunit rRNA [13]. Group I introns have also been found to interrupt a large variety of tRNAs, mRNAs and rRNAs. They all have a common secondary structure and a common splicing pathway [14]. The secondary structure model is defined by conserved structural (P1–P10) and sequence elements (P, Q, R and S; Fig. 1a), which are essential for catalysis [15–24]. Ribozyme sequence specificity is defined by the interactions between the 5′-region of the intron and the two exons (the P1 and P10 domains; Fig. 1a). The intron sequence, which defines the cleavage site and interacts with the 5′-exon, is called the IGS (internal guide sequence). The reaction at the 5′-splice site has been well characterized, with cleavage occurring downstream of a conserved U–G pair. Similarly, 3′-splicing occurs immediately 3′ of a universally conserved guanosine (Fig. 1a) [14]. Figure 1 Open in new tabDownload slide Group I introns. a: Schematic representation of the consensus secondary structure of group I introns. P1–P9, conserved double-stranded elements. A dashed double-head arrow indicates P10 interaction between the exon 2 and the 5′-region of the intron (shown in gray). The indicated phylogenetically conserved sequence elements P, Q, R and S, are represented by the most common sequences. A thick line indicates the IGS. Exon sequences are depicted in boxes. Arrows indicate the 5′- and 3′-splice sites. Figure adapted from Cech [299]. b: Main stages of the group I self-splicing mechanism. Solid lines denote the intron and exons are represented by boxes. Exogenous guanosine, essential for catalysis, is shown encircled. Figure adapted from Cech [300]. Figure 1 Open in new tabDownload slide Group I introns. a: Schematic representation of the consensus secondary structure of group I introns. P1–P9, conserved double-stranded elements. A dashed double-head arrow indicates P10 interaction between the exon 2 and the 5′-region of the intron (shown in gray). The indicated phylogenetically conserved sequence elements P, Q, R and S, are represented by the most common sequences. A thick line indicates the IGS. Exon sequences are depicted in boxes. Arrows indicate the 5′- and 3′-splice sites. Figure adapted from Cech [299]. b: Main stages of the group I self-splicing mechanism. Solid lines denote the intron and exons are represented by boxes. Exogenous guanosine, essential for catalysis, is shown encircled. Figure adapted from Cech [300]. The self-splicing reaction proceeds via a two-step transesterification mechanism initiated by nucleophilic attack by the 3′-OH group of an exogenous guanosine nucleotide at the phosphodiester bond between the last nucleotide of exon 1 and the 5′-end nucleotide of the intron (5′-splice site, Fig. 1b). This results in the covalent linkage of the guanosine to the 5′-end of the intron and the release of the 3′-OH free-exon 1, which remains bound to the intron by base pairing. In the second step, the phosphodiester bond between the intron and exon 2 (3′-splice site) is attacked by the 3′-OH group of exon 1, resulting in excision of the intron with the guanosine at its 5′-end, and the release of the ligated exons. The reaction is completely reversible, so the intron is able to integrate between both exons [25]. The Tetrahymena group I intron has been re-engineered to yield a true catalyst that cleaves exogenous substrate molecules in trans. For reactions to occur, substrates must contain the complementary sequence of the ribozyme IGS and maintain the essential U–G base-pair that defines the cleavage site [26]. No high-resolution crystal structure of an entire group I ribozyme with bound substrates has yet been determined. However, the crystal structures of different truncated forms of the Tetrahymena intron reveal that the RNA molecule folds into a globular structure with an interior relatively inaccessible to solvents [27, 28]. The main strategies developed for the application of group I introns as specific gene silencers are described in Section 3.1.3. 2.1.2 Group II introns Group II introns are large catalytic RNA molecules that fold into a compact structure essential in RNA catalyzed self-splicing and intron mobility reactions. They are commonly found in bacteria and the organellar genes of plants, fungi and yeast (reviewed by [29, 30]). They are mainly present in mRNAs, but they also occur in tRNA and rRNA genes. The secondary structure is represented by six helical domains (I–VI) emerging from a central domain (Fig. 2a). This model was derived from extensive phylogenetic and mutational analyses [30]. It has been reported that only domains I and V are indispensable for catalysis [31]. Further, domain V has been described as the reaction center [32, 33]. Two major subgroups of group II introns, IIA and IIB, have been defined, mainly based on their tertiary interactions. Figure 2 Open in new tabDownload slide Group II introns. a: Two-dimensional structural model of group II introns. The main structural domains of the intron are represented (I–VI), and the conserved adenosine implicated in the first nucleophilic attack is shown within domain VI. IBSs and EBSs are depicted in black boxes, as well as ε and ε′ sequences. Exons are depicted in boxes while a solid line shows the intron. Figure adapted from Carola and Eckstein [301]. b: Pathways for the group II intron self-splicing reaction. Intron is depicted as a solid line and exon sequences are shown by boxes. Figure adapted from Pyle [29]. Figure 2 Open in new tabDownload slide Group II introns. a: Two-dimensional structural model of group II introns. The main structural domains of the intron are represented (I–VI), and the conserved adenosine implicated in the first nucleophilic attack is shown within domain VI. IBSs and EBSs are depicted in black boxes, as well as ε and ε′ sequences. Exons are depicted in boxes while a solid line shows the intron. Figure adapted from Carola and Eckstein [301]. b: Pathways for the group II intron self-splicing reaction. Intron is depicted as a solid line and exon sequences are shown by boxes. Figure adapted from Pyle [29]. The splicing of group II introns proceeds via two transesterification steps (reviewed by [29]). The first involves nucleophilic attack at the 5′-splice site by the 2′-OH group of a conserved adenosine located in domain VI (Fig. 2b) [33]. As in nuclear mRNA introns, the reaction yields a lariat structure where the 5′-end of the intron is ligated to the adenosine (branch site) by a 2′,5′-phosphodiester bond. The second reaction occurs by the free 3′-OH of exon 1 attacking the 3′-splice site, resulting in the ligation of the two exons and the release of the intron lariat [34]. Some group II ribozymes are thought to be able to insert themselves into double-stranded DNA by reverse splicing and transcription [35], thus promoting the mobility of these elements. Group II introns can also catalyze trans reactions. The substrate sequence recognition can be quite long (>13 nt), and reactions can proceed with a very high degree of sequence specificity [36]. Recognition of the 5′-exon is achieved by the interaction of IBS1 and IBS2 sequences (intron binding sequences in the 3′-end of the exon) with the specific sequences EBS1 and EBS2 (exon binding sequences) found in domain I of the intron (Fig. 2a). Further interaction of ε and ε′ sequences helps to define the 5′-splice site. Reactions of different nature catalyzed by group II introns have been described. Group II ribozymes can catalyze the cleavage of ligated RNA precursors, ligate RNA to DNA and cleave single-stranded DNA substrates [37]. Moreover, the ability of the group II introns to insert themselves into double-stranded DNA opens up the possibility of using these ribozymes to target and inactivate specific pathogen genes [38]. 2.2 RNase P RNase P is a key enzyme in the biosynthesis of tRNAs. It is found in all cells and organelles that carry out tRNA synthesis [39, 40]. It is an RNA processing endonuclease that specifically cleaves the tRNA precursors, releasing 5′-sequences and mature tRNAs (Fig. 3). All known RNase P enzymes are ribonucleoproteins (RNPs) that contain an RNA subunit essential for catalysis, with the possible exception of RNase P from some plant chloroplasts and Trypanosomatid mitochondria [41, 42]. Although the protein component is needed for the in vivo activity of the enzyme [43], it has been shown that, at least for several eubacterial RNase P enzymes, the RNA component is the catalytic subunit. This was first described by Altman and co-workers, who demonstrated that the RNA component from Escherichia coli RNase P (M1 RNA) was sufficient to catalyze the cleavage reaction in vitro [44]. M1 RNA is responsible for substrate binding and cleavage, while the protein subunit diminishes the ionic strength required for optimal activity and increases the reaction rate by facilitating product release after cleavage [45]. The ribozyme is thought to be a metalloenzyme [46, 47]. The reaction is a one-step cleavage that generates the very characteristic 5′-phosphate (mature tRNAs) and 3′-hydroxyl (5′-precursor sequences) products (Fig. 3a). Figure 3 Open in new tabDownload slide RNase P cleavage. a: Schematic representation of the cleavage reaction of the natural RNA substrate by the RNase P enzyme. In this reaction the 5′-end of the pre-tRNA is eliminated to generate mature tRNAs. The conserved triplet CCA at the 3′-end of the substrate RNA, needed for catalysis, is shown. b: Structure model of the minimal target RNA–EGS complex cleavable by bacterial RNase P, only a double-stranded domain carrying the CCA triplet is needed for cleavage. c: Target RNA-3/4EGS complex that can be recognized as substrate by eukaryotic RNase P. d: Representation of a minimized 3/4EGS-target complex. In all cases the arrow indicates the cleavage site. A thick line indicates the substrate RNA. Figure 3 Open in new tabDownload slide RNase P cleavage. a: Schematic representation of the cleavage reaction of the natural RNA substrate by the RNase P enzyme. In this reaction the 5′-end of the pre-tRNA is eliminated to generate mature tRNAs. The conserved triplet CCA at the 3′-end of the substrate RNA, needed for catalysis, is shown. b: Structure model of the minimal target RNA–EGS complex cleavable by bacterial RNase P, only a double-stranded domain carrying the CCA triplet is needed for cleavage. c: Target RNA-3/4EGS complex that can be recognized as substrate by eukaryotic RNase P. d: Representation of a minimized 3/4EGS-target complex. In all cases the arrow indicates the cleavage site. A thick line indicates the substrate RNA. The RNA component of RNase P from bacteria varies in length from 350 to 410 nucleotides (nt) [48]. Despite sequence conservation among the known bacterial examples being limited to short regions, a consensus secondary structure has been defined [49]. This came from extensive phylogenetic covariation analysis of related sequences [48, 50]. Typically, it comprises two domains containing the substrate-recognition site and the ribozyme active site. RNase P has long been thought useful as a gene silencer since it is ubiquitous. Substrate recognition is based on structural features more than on sequence features. Altman and co-workers showed that the substrate requirements of E. coli RNase P are limited to a short double-stranded region with CCA at the 3′-end (Fig. 3b) [51, 52]. This is the basis of the gene-silencing strategy developed for the RNase P ribozyme. It is proposed that E. coli RNase P can target any RNA molecule as long as the external sequence-specific RNA molecule (external guide sequence, EGS) is present. The target RNA and the EGS would together form a double-stranded domain carrying the CCA triplet at the 3′-end of the EGS. This structure mimics the natural substrates of RNase P and can therefore be recognized by the enzyme (reviewed by [53]). In human cells, the RNase P complex is longer and includes multiple protein components in addition to the RNA [54–56]. The human RNase P RNA has not been shown to be catalytically active in the absence of these protein components. In contrast to bacterial RNase P, eukaryotic RNase P is unable to cleave the model complex defined for E. coli RNase P. Initial studies on recognition of bipartite substrates by human RNase P showed that cleavage occurs when three-quarters of the tRNA molecule is presented as the EGS (Fig. 3c) [57]. This EGS, called 3/4EGS, forms a sequence- and structure-specific complex with the corresponding target RNA. Further studies demonstrated that the 3/4EGS can be minimized without drastically altering recognition and cleavage of the bimolecular substrate by human RNase P (Fig. 3d) [57, 58]. In this modified design, the EGS is about 30 nt in length and is complementary to 11 nt in the target RNA. The main advantage of this strategy is that the cellular environment is optimal for RNase P catalysis, and it should be the environment in which maximum catalytic efficiency is reached. 2.3 Small catalytic RNAs This class of ribozyme comprises several types of catalytic RNAs with common features. First, they are small compared to the previously described ribozymes. Second, they are all naturally involved in the replication process of RNA genomes in which they are contained. Finally, in all cases, the reaction catalyzed by these ribozymes proceeds via transesterification chemistry that generates 5-hydroxyl and 2′,3′-cyclic phosphate termini [59]. However each group is defined by a characteristic secondary structure. 2.3.1 Hammerhead ribozyme The hammerhead ribozyme was first discovered as a catalytic motif in different plant pathogen RNAs. All hammerhead ribozymes share a characteristic secondary structure (Fig. 4) [60]. Currently, this structure is known in several plant pathogen RNAs: the genomic RNA of three different plant viroids, [6, 61, 62] nine satellite RNAs [60, 63–66], a circular RNA from cherry [67], and a retroviroid-like element of carnation plants [68]. In addition, three active hammerhead domains isolated from animal RNAs have also been described: a transcript from a satellite DNA of newts [69], the RNA encoded in Schistosoma satellite DNA [70], and in a DNA satellite from Dolichopoda cave crickets [71]. All of them are involved in the processing of long multimeric transcripts into monomer-sized molecules. The plant-derived ones play an essential role in the in vivo replication process of RNA genomes in which they are contained. Replication of these RNAs occurs by the rolling circle mechanism [65, 72, 73]. During this process, multimeric products are generated which have to be converted into genome-length strands. It is known, for at least for 14 of these RNAs, that this involves a self-cleavage reaction catalyzed by the hammerhead domain. Figure 4 Open in new tabDownload slide Trans-cleaving hammerhead ribozyme. Secondary structure model of the hammerhead ribozyme–substrate complex. Important nucleotides for catalytic activity and structural domains helices I to III are shown. Ribozyme nucleotides are in uppercase letters; substrate nucleotides are in lowercase letters. The arrow indicates the cleavage site. Nucleotides are numbered as described in [302]. • represents any nucleotide; Y represents C or U; R represents A or G; h is A, C or U. Figure 4 Open in new tabDownload slide Trans-cleaving hammerhead ribozyme. Secondary structure model of the hammerhead ribozyme–substrate complex. Important nucleotides for catalytic activity and structural domains helices I to III are shown. Ribozyme nucleotides are in uppercase letters; substrate nucleotides are in lowercase letters. The arrow indicates the cleavage site. Nucleotides are numbered as described in [302]. • represents any nucleotide; Y represents C or U; R represents A or G; h is A, C or U. Molecular modeling and kinetic analysis of the hammerhead cleavage reaction in the presence of monovalent or divalent salts support the idea that divalent metal ions are not essential for the catalytic step, although they do stabilize the structure of active ribozymes [74–77]. The minimal motif that supports catalytic activity was defined by deletion assays [6, 12, 60, 78, 79]. It has been divided into two molecules, making trans-cleavage reactions possible [10, 80]. The hammerhead motif most commonly used is a 35-nt-long RNA molecule, but this varies depending on the length of the substrate binding arms (Fig. 4). The hammerhead ribozyme–substrate complex is comprised of an intramolecular helix (helix II) and two intermolecular helices generated after the substrate interaction (helix I and III). Single-stranded regions are highly conserved and contain most of the important nucleotides for optimum catalytic activity [81]. However, substrate-binding arms can be changed to modify ribozyme specificity. Helix I shows no strict sequence requirements. Nevertheless, it has been shown that the helix I region close to the catalytic core has some influence on global ribozyme structure [82]. The nucleotide sequence of this region defines the angle between helices II and III, contributing to active conformer formation (Fig. 4). The consensus sequence for the cleavage site has been established as 5′-NHH↓ (N=any nucleotide and H=A, C or U; ↓=the cleavage site) [83]. The three-dimensional structure of the hammerhead ribozyme was established by two different approaches: by the X-ray diffraction spectrum of the ribozyme when co-crystallized with a DNA-substrate molecule [84, 85], and by FRET techniques [86]. The data shows that the ribozyme adopts a ‘Y’ shape in which helices II and III are co-linearly stacked with helix I adjacent to helix II. The hammerhead ribozyme is probably the most widely studied ribozyme. It is also the most commonly used for gene inactivation assays due to its small size and catalytic efficiency. 2.3.2 Hairpin ribozyme The first naturally occurring hairpin motif was identified in the negative strand of the satellite RNA associated with the tobacco ringspot virus ([-]sTRSV) [7, 87]. Only two additional natural hairpin ribozymes have been identified, both from plant satellite viruses [88]. The hairpin catalytic motif is involved in vivo in the replication process of the satellite RNAs in which they are found. It processes the multimeric products generated during rolling circle replication into genome length monomers, as described for hammerhead, but the hairpin motif also catalyzes the ligation of both ends, generating circular RNA forms [89]. The minimal sequences required for catalytic activity were defined by a combination of mutagenesis, insertions and deletions [7, 80, 87]. Further, the catalytic domain was engineered to support trans-cleavage reactions (Fig. 5a) [7, 87]. The hairpin catalyzes the reversible cleavage of a second RNA molecule (substrate) using a transesterification mechanism. The reaction is metal-ion independent, but metal ions can play a role in structure stabilization [90–92]. The hairpin ribozyme is also an effective RNA ligase [93]. Figure 5 Open in new tabDownload slide Trans-cleaving hairpin ribozyme. a: Secondary structure model of the [-]TRSV hairpin ribozyme–substrate complex. Ribozyme nucleotides are numbered 1–50, and substrate nucleotides are numbered −5 to +9. H1–H4 and J1/2–J4/3 are double- and single-stranded regions respectively, as described by [94, 95]. A and B represent the two defined domains of the ribozyme–substrate complex. An arrow indicates the cleavage site. b: Sequence requirements for the hairpin ribozyme–substrate complex. N represents any nucleotide; Y represents C or U; R represents A or G; B represents C, G or U; V represents A, C or G. Important nucleotides are shown in bold letters. Substrate sequences of J2/1 region more efficiently cleaved are shown in box (a). The optimizing U39 to C substitution is also included. Figure 5 Open in new tabDownload slide Trans-cleaving hairpin ribozyme. a: Secondary structure model of the [-]TRSV hairpin ribozyme–substrate complex. Ribozyme nucleotides are numbered 1–50, and substrate nucleotides are numbered −5 to +9. H1–H4 and J1/2–J4/3 are double- and single-stranded regions respectively, as described by [94, 95]. A and B represent the two defined domains of the ribozyme–substrate complex. An arrow indicates the cleavage site. b: Sequence requirements for the hairpin ribozyme–substrate complex. N represents any nucleotide; Y represents C or U; R represents A or G; B represents C, G or U; V represents A, C or G. Important nucleotides are shown in bold letters. Substrate sequences of J2/1 region more efficiently cleaved are shown in box (a). The optimizing U39 to C substitution is also included. The most widely used hairpin model is that described by Hampel and Triz (Fig. 5a) [7]. The ribozyme is a 50-nt-long RNA that recognizes and cleaves an external 14-nt-long substrate RNA. The secondary structure of the ribozyme–substrate complex was established through mutational and in vitro selection analysis [94–96]. It is formed by two, well defined, independently folding domains, A and B (Fig. 5) [97]. Each has two helical regions separated by an internal loop. Domain A is formed by both the ribozyme and the substrate RNA molecules and consists of the two intermolecular helices (1 and 2) and the internal loop A. Domain B is entirely composed of the ribozyme sequences (helices 3 and 4 and loop B). After formation of the ribozyme–substrate complex, an essential conformational change must occur. Domain A folds upon domain B and the nucleotides of both loops interact. This process, known as docking, is essential for the activity of the hairpin ribozyme [92, 98–100]. The crystal structure of the hairpin ribozyme was recently obtained [101]. It shows a solvent-excluded cleft in the ribozyme active site. This crystal structure also shows that an active-site adenosine might play an important role in a general acid–base catalysis, but nucleotide analogue interference experiments have not provided evidence for this [102]. However, the proposed metal ion-independent mechanism is consistent with the lack of divalent metal ions requirements for hairpin ribozyme cleavage [90, 91, 103]. Most of the nucleotides important for catalytic activity are contained within the impaired regions. There are almost no sequence restrictions for bases involved in the formation of the helices as long as base pairing is achieved (Fig. 5b) [95]. Substrate recognition and binding by the ribozyme involves the formation of the two intermolecular helices – helix 1 and 2- (of 6 and 4 bp respectively) [104, 105]. Therefore, the specificity of the ribozyme can be altered to target practically any RNA molecule of interest. Similarly, substrate regions that interact with the ribozyme do not show high sequence requirements. However, a stable association between the two molecules is required, especially for those nucleotides close to loop A (and therefore close to the cleavage site). The single-stranded region that contains the cleaved bond (J2/1 region in Fig. 5b) shows strict sequence requirements. A G at position +1 is essential for the cleavage reaction [106]. Nevertheless, it has recently been reported that mutations of G+1 can be tolerated in the presence of the compensatory changes at ribozyme nucleotide C25 [107]. This suggests a base-base interaction occurs – G+1/C25 – after docking. Although it has not been possible to reach a general consensus as to the substrate J2/1 sequence requirements that would allow efficient cleavage by the hairpin ribozyme, detailed information exists on the cleavage efficiency of all the possible sequence combinations of the J2/1 region nucleotides, except for G+1 [108]. Only 40 variants are processed, and the optimal substrates have been defined [108]. Many efforts have been made to improve the catalytic performance of the hairpin ribozyme by sequence or structural modifications of the minimal active ribozyme. A positive effect can be achieved by substitution of the U39 by C [94, 109, 110] or by the stabilization of helix 4 [96, 110–112]. 2.3.3 HDV ribozyme The HDV is an infectious agent that exits as a satellite RNA of the hepatitis B virus [113]. Its genome is a single-stranded circular RNA of 1700 nt. It is thought to replicate by the double rolling-circle mechanism, which requires self-cleavage by closely related versions of a catalytic domain contained within the genomic and antigenomic RNAs [114–116]. The two ribozymes are similar in sequence and structure, though the proposed common secondary structure differs from those of other, small, catalytic RNAs (Fig. 6). The minimal self-cleavage domain has been defined as an 85-nt-long fragment, with 1 nt 5′ of the cleavage site (−1 in Fig. 6) and 84 nt 3′ of the cleavage site [8, 117]. However, recent studies suggest that although positions −2 to −4 are dispensable, they could be involved in ribozyme specificity [118]. The proposed secondary structure has four double-stranded regions (P1–P4; Fig. 6). P1 and P2 would form a pseudoknot structure; P3 and P4 are stem–loop motifs. In this model, the P1 helix defines the cleavage site. Figure 6 Open in new tabDownload slide HDV ribozymes. Secondary structure representation of the genomic an antigenomic HDV ribozymes. Dotted lines in 3′ and 5′ represent additional sequences not required for self-cleavage. Continuous lines connect different structural elements, they do not represent omitted sequences. P1–4, double-stranded regions. Cleavage sites are indicated by arrows. Figure adapted from Perrota and Been [8]. Figure 6 Open in new tabDownload slide HDV ribozymes. Secondary structure representation of the genomic an antigenomic HDV ribozymes. Dotted lines in 3′ and 5′ represent additional sequences not required for self-cleavage. Continuous lines connect different structural elements, they do not represent omitted sequences. P1–4, double-stranded regions. Cleavage sites are indicated by arrows. Figure adapted from Perrota and Been [8]. As described for the hammerhead and hairpin ribozymes, the catalytic activity of HDV is more efficient in the presence of divalent cations. Recent progress has been made in understanding the catalytic mechanism. Of great importance has been the resolution of the crystal structure of the genomic ribozyme [119, 120]. The structure shows a deep active site cleft in which well-ordered metal ions were not observed. Moreover, it shows a specific cytosine (C75, Fig. 6) positioned to act as a general acid–base catalyst [121–123]. Minimal delta self-cleavage RNA can be separated into two molecules in order to develop trans systems, generating delta ribozymes that can catalyze the successive cleavage of several molecules of substrate [124]. For designing these trans-active molecules, the 5′-strand of the P1 helix was eliminated from the catalytic domain. This sequence is provided by a second RNA molecule acting as substrate. The sequence specificity is defined by the 3′-P1 sequence. A circular variant of this trans-acting ribozyme has been developed [125]. The cleavage rate of the circular HDV-like ribozyme is comparable to that of the linear HDV ribozymes containing the same core sequence, but it shows greater resistance to nuclease activity. HDV ribozymes are very stable ex vivo, with an estimated half-life of over 100 h. The P2 stem has a critical role in this stability [126]. HDV ribozyme appears to be well adapted to the human cell environment and could be ideal for the development of a gene-inactivation system. However, despite the theoretical advantages of the ex vivo use of these ribozymes, few ex vivo trans-cleavage experiments have been performed [127]. 2.3.4 VS ribozyme The VS ribozyme was isolated from the mitochondria of the natural Varkud-1c strain of Neurospora[128]. This catalytic motif is responsible of the self-cleavage and ligation of the non-encoding VS RNA [129], which is transcribed from the VS plasmid DNA [130]. The catalytic activity has been restricted to a 154-nt-long fragment, retaining 1 nt 5′ of the cleavage site and 153 nt at its 3′[131]. A further reduction of this domain has been achieved by deletions of internal regions within the different structural domains, limiting the catalytic fragment to 126 nt [132]. The proposed secondary structure is composed of six defined domains (I–VI) and implies an essential interaction between the nucleotides of loops I and V (Fig. 7). Further, structural rearrangements are required for ribozyme catalytic activity [133, 134]. Figure 7 Open in new tabDownload slide VS ribozyme. Secondary structure of the VS RNA ribozyme. Structural domains I–VI are represented. Numbering is that of the full-length VS RNA. Nucleotides implicated in the interaction between loops I and V are indicated. An arrow shows the cleavage site. Figure adapted from Beattie et al. [303]. Figure 7 Open in new tabDownload slide VS ribozyme. Secondary structure of the VS RNA ribozyme. Structural domains I–VI are represented. Numbering is that of the full-length VS RNA. Nucleotides implicated in the interaction between loops I and V are indicated. An arrow shows the cleavage site. Figure adapted from Beattie et al. [303]. As described for the other catalytic motifs, the minimal self-cleaving RNA domain can be separated into two molecules in order to develop trans-acting ribozymes [135]. For designing these ribozymes, the domain I is provided by a separated RNA molecule, which is efficiently cleaved by a 144-nt-long VS catalytic domain. The global structure of the trans-acting ribozyme has been recently solved [136]. These authors proposed that the substrate, a stem–loop motif, interacts with the ribozyme by docking between domains II and VI, while making a tertiary loop–loop contact with the domain V. The substrate recognition by the ribozyme does not imply the formation of stable intermolecular helical regions between ribozyme and substrate, as it has been described for other ribozymes (e.g., hammerhead and hairpin). In contrast, the VS ribozyme recognizes a stable stem–loop, and the recognition is determined by stem–loop interactions [135]. Therefore it has been proposed that the VS ribozyme might be appropriate to target structured RNA substrates. 3 Ribozyme-based therapeutics Ribozymes are strong candidates for development as therapeutic agents. Strategies are already being designed that involve ribozymes fighting viral (e.g., AIDS, viral hepatitis) and cellular diseases (e.g., cancer, diabetes, rheumatoid arthritis) – maladies against which conventional therapy has had little success. The successful use of ribozymes as therapeutic agents depends upon many factors, e.g., the relative amounts of active ribozymes in cells, their co-localization with target RNAs, the structural features of transcripts that influence accessibility to specific target sites, catalytic efficiency, the interaction of target RNAs with proteins, and the intracellular stability of targeted RNAs and ribozymes. The solutions to some of the problems of ribozyme therapy are quite simple and require no redesigning of the catalytic RNA domains. For example, the relative amounts of ribozymes inside cells and their localization can be controlled by using different promoters to drive ribozyme synthesis [127]. Several authors have described the use of expression systems in which different ribozymes with different specificities are encoded by the same cassette [137]. Moreover, the catalytic performance of ribozymes can be improved by simple sequence modifications [109, 112], and greater RNA stability can be achieved by chemical modification [138]. Other desirable characteristics such as enhanced substrate specificity, improved accessibility of specific target sequences within long, folded RNA molecules, increased turnover rates and allosteric regulation, have been obtained by modifying hammerhead and hairpin ribozymes. 3.1 Plain ribozymes 3.1.1 Small ribozymes The development of specific gene silencing tools – and therefore of ribozyme therapeutic strategies [139, 140]– has heavily involved the hairpin- and hammerhead-type ribozymes, members of the group of small catalytic RNAs. A huge number of ex-vivo experiments aimed at achieving specific gene inactivation have been performed with trans-acting hairpin or hammerhead ribozymes. Further, both types are currently being tested in gene therapy clinical trials. Hepatitis B virus RNAs (HBV) have been targeted by hammerhead and hairpin ribozymes. Inhibition of viral replication has mainly been achieved by targeting HBx mRNA [141, 142]. The HBx gene encodes a transcriptional activator protein involved in both viral replication and the development of virus-associated disease (hepatitis, cirrhosis and hepatocellular carcinoma). A reduction in protein level – and consequently in trans-activating capacity – has been achieved by targeting this gene at the RNA level with hammerhead ribozymes. Moreover, a reduction in HbsAg and HbeAg surface antigens is also observed, suggesting that virus replication is inhibited [141, 142]. In a human liver cell line, 80% inhibition of HBV replication was achieved by targeting the polymerase and X protein mRNAs with hairpin ribozymes [143]. The inhibition of virus replication has also been accomplished by targeting pregenomic RNA [144]. Hepatitis C virus infections are particularly suited to ribozyme-mediated gene therapy since the viral genome is exclusively present as RNA. The highly conserved 5′-untranslated region (UTR) and the coding sequence for the nucleocapsid have been the main targets. The use of specific hammerhead ribozymes has notably reduced HCV RNA from hepatocytes of chronic HCV-infected patients [145]. Such a significant reduction in viral load is of great importance in improving the quality of life of patients with chronic infection. Moreover, nuclease-resistant hammerhead ribozymes have been shown to significantly inhibit replication of an HCV-poliovirus chimera by targeting different sites in the 5′-UTR [146]. Other experiments performed with hairpin catalytic motifs have demonstrated that ribozymes are responsible for cellular resistance to infection by retroviral particles containing HCV target sequences [147]. A clinical trial using modified hammerhead ribozymes against HCV RNAs is underway [148]. HIV-1 RNAs are favorite targets in ribozyme inhibition assays with hairpin and hammerhead ribozymes. The inactivation of the RNAs of the tat, env, gag or pol genes has been the focus of many tests [149–157]. The highly conserved 5′-LTR region and the packing signal ψ have also been used widely [158–163]. Many of the tested ribozymes have demonstrated efficacy at inhibiting HIV replication in cultured cells, and spectacular reductions in p24 production and viral RNA levels have been achieved. A very elegant strategy consists of introducing the ribozymes into the virion, requiring they be linked to tRNAlys (the primer used during lentivirus replication) [164]. Other anti-HIV ribozyme strategies have been aimed at trying to block viral entry to cells. For this purpose chemokine receptor CCR5 mRNA has been targeted [165–167], resulting in significant resistance to viral infection with only a 20% downregulation of the receptor [166]. Retroviral vectors are being used in clinical trials in which anti-HIV hammerhead [168, 169] and hairpin ribozyme [170] expression cassettes are introduced into CD4+ lymphocytes or CD34+ hematopoietic precursors (ex vivo) taken from HIV-1-infected patients and their identical, non-infected twins. Once the cells are transduced they are infused into the patient and the survival of ribozyme-containing cells monitored. Initial results suggest that the transfer of ribozyme-encoding genes to HIV-1-infected individuals is well tolerated, and that transduced cells persist in the patient [170]. Unfortunately, it seems that they fall below detection levels 1 year after infusion [171]. The selective destruction of the RNAs of mumps virus [172], influenza virus [173], alphavirus [174] and human papillomavirus type 16 [175] has also been accomplished with these small ribozymes. Other targets that have attracted attention are those involved in the induction or progression of tumors [176–180] and genes important in cancer therapy (such as MDR-1– multidrug resistance). A hammerhead-mediated reversion of the multiple-drug-resistant phenotype, accompanied by restoration of sensitivity to chemotherapeutic agents, has been achieved by inactivating this gene in cancer cells [181–183]. Two clinical trials are in progress to evaluate the potential of chemically modified hammerhead ribozymes to fight cancer. The first, a phase II trial designed to test therapeutic efficacy in breast and colorectal cancer [171], involves a hammerhead ribozyme that targets flt-1 mRNA. This encodes the high affinity receptor for vascular endothelial growth factor, an angiogenic protein. Daily intravenous or subcutaneous delivery of this ribozyme is well tolerated and plasma levels are maintained for prolonged periods after subcutaneous delivery [148]. In the second trial, the target is human epidermal growth factor receptor type 2 mRNA, which is overexpressed in many breast cancers [138]. RNAs involved in other cellular diseases, for example Alzheimer's disease [184], rheumatoid arthritis [185] and diabetes [186], have also been targeted by ribozymes. 3.1.2 RNase P RNase P has been used for the selective inhibition of pathogenic RNAs. Two different procedures have been developed based on RNase P catalytic activity. The first takes advantage of endogenous RNase P and requires only the delivery of an EGS [41, 187]. The second is based on the use of a modified external catalytic RNA component of a heterologous RNase P (e.g., E. coli M1 RNA), which carries the guide sequence (GS) covalently linked at its 3′-end [188]. The first strategy has been used more frequently since the cellular environment provides the optimal reaction conditions for endogenous RNase P. Further, only a short RNA molecule (EGS) has to be delivered into the cell. This technology has been shown effective at inactivating several viral genes. Liu and co-workers designed an EGS to target the mRNA encoding the thymidine kinase (TK) of herpes simplex virus 1 (HSV-1), and achieved an 80% reduction in TK mRNA and protein expression in human cells steadily expressing the EGS [189]. HIV-1 mRNAs have also been targeted by human RNase P. In this case, the EGS was designed to target the highly conserved HIV-U5 region and was introduced into a CD4+ T cell line which was later infected with HIV-1. Cells treated with U5-EGS maintained their CD4+ expression and lack of HIV p24 antigen production, suggesting that HIV-1 replication might be effectively inhibited by this ‘protective’ U5-EGS [190]. Similarly, other authors have successfully inhibited influenza virus production in mouse cells by using a specific EGS [191]. With this virus, better inhibition can be achieved by simultaneously targeting two different sites within the same target RNA [191]. The possibility of using a DNA-based EGS instead of an RNA-based EGS has also been demonstrated. This strategy was used by Liu and co-workers to target the mRNA encoding the protease of human cytomegalovirus (HCMV) [192]. EGSs were chemically synthesized and exogenously administered into HCMV-infected human foreskin fibroblasts. A reduction of about 80–90% in the protease level and a 300-fold drop in HCMV replication were observed [192]. In all the cases described above, the EGSs were designed to bind the target RNA and resemble the natural tRNA structure. However, in vitro-selected EGSs can be more efficient for RNase P-mediated gene inactivation [193]. An EGS selected for the TK mRNA of HSV-1 was 35 times more active than the EGS derived from a natural tRNA sequence, achieving a 95% reduction in TK mRNA in HSV-1-infected cells. This selected EGS, when in complex with the target mRNA, resembles a portion of the tRNA structure and shows enhanced binding affinity for the target RNA. The second strategy for using RNase P based-ribozymes involves a modified M1 RNA converted into an endoribonuclease that provides the corresponding GS linked to its 3′-end (M1GS ribozyme) [188, 194]. M1GS ribozyme has been described to cleave the TK mRNA of HSV-1 both in vitro and in mouse cells ex vivo [194]. Similarly, Liu and co-workers reported the use of a M1GS RNA for targeting the mRNA encoding the human HSV-1 major transcription activator, ICP4 [195]. A reduction of more than 80% in the expression of ICP4, and about a 1000-fold drop in viral growth, were observed in cells that stably expressed the ribozyme. This catalytic M1 RNA has also been used for targeting cellular RNAs – specifically the bcr-abl oncogenic messengers [196]. M1GS RNA was designed to recognize the oncogenic messenger at its fusion point. The effectiveness and specificity of the ribozyme was tested in vitro and in mammalian cell models. Significant reductions in Bcr-abl mRNA levels and the prevention of bcr-abl oncogene functions were observed. M1 RNA has also been subjected to in vitro selection methods in order to render improved M1 ribozymes. These engineered RNase P ribozymes have been successfully employed to target human HSV-1 [197] and HCMV [198, 199], both in vitro and in mammalian cells. The molecular study of the selected variants suggests that point mutations can both increase the rate of chemical cleavage and enhance substrate binding [198]. Interestingly, these results show there is a possibility of obtaining highly effective RNase P ribozymes with attractive therapeutic features. Finally, it has recently been reported that human RNase P can cleave HCV RNA in the absence of EGSs [200]. These authors report that HCV genomic RNA is specifically and efficiently cleaved by purified RNase P from HeLa cells. 3.1.3 Self-splicing introns Apart from therapeutic strategies aimed at destroying harmful RNA molecules, a technology based on the use of group I ribozymes is being pursued that might allow the repair of mutant RNAs. Group I ribozymes can be designed to cut upstream of a nonsense or a missense mutation and splice in a corrected transcript to restore the correct genetic information (Fig. 8) [201]. The ribozyme carries an exogenous exon-like attached to its 3′-end, the 5′-end of which corresponds to the nucleotide immediately 3′ of the cleavage site and extends to the end of the sequence. This exon provides the corrected sequence and is spliced in trans to the endogenous RNA. The ribozyme first cleaves out the RNA fragment containing the mutation and then catalyzes the ligation of the newly formed 3′-end to the exogenous exon. This reaction, which generates a recombinant transcript, has been generically called trans-splicing. This strategy could also be used to convert RNAs encoding harmful proteins into innocuous forms. Figure 8 Open in new tabDownload slide Trans-splicing by group I ribozymes. Scheme of the proposed strategy for reparation of an aberrant RNA by a group I ribozyme. Trans-splicing group I ribozymes recognize a mutant RNA upstream of a mutation site. The recognition is performed through the IGS present in the 5′-end of the ribozyme. The mutant RNA is cleaved up-stream of the mutation and the 3′-fragment is replaced by the sequences carried as a 3′ exon-like fragment by the ribozyme, generating a correct transcript. Figure adapted from Sullenger and Cech [201]. Figure 8 Open in new tabDownload slide Trans-splicing by group I ribozymes. Scheme of the proposed strategy for reparation of an aberrant RNA by a group I ribozyme. Trans-splicing group I ribozymes recognize a mutant RNA upstream of a mutation site. The recognition is performed through the IGS present in the 5′-end of the ribozyme. The mutant RNA is cleaved up-stream of the mutation and the 3′-fragment is replaced by the sequences carried as a 3′ exon-like fragment by the ribozyme, generating a correct transcript. Figure adapted from Sullenger and Cech [201]. RNA message correction by group I ribozymes was first attempted by Sullenger and Cech [201]. They successfully repaired a truncated form of the lacZ mRNA both in vitro and in E. coli cells, obtaining functional β-galactosidase. Subsequently, they demonstrated that targeted trans-splicing proceeded efficiently in murine fibroblasts with high fidelity [202]. In fact, they achieved 50% corrected lacZ substrate in this system [203]. However, they also observed a relatively low specificity of the ribozyme-catalyzed reaction. Trans-splicing ribozymes reacted with the targeted substrate RNAs but also with ‘unintended’ transcripts. This problem has been partially overcome by the extension of the IGS (and therefore of the P1 interaction region) and by the addition of an antisense domain complementary to the target RNA [204]. Other authors have focused on therapeutically important targets, such as trinucleotide repeated expansions in muscular dystrophy protein kinase transcripts [205], sickle cell β-globin mRNAs [206], and, more recently, mutant p53 transcripts [207]. Trans-splicing can also be used to create functional chimeric mRNAs to be expressed exclusively in those cells that contain a chosen mRNA target. The RNA sequences provided by trans-splicing ribozymes would be expressed solely as the resultant chimeric mRNA. This strategy could be used to genetically mark or kill malignant or otherwise harmful cells, including those infected by viruses [208]. Group II introns are a second type of self-splicing intron, which can function as mobile genetic elements. Their mobility is mediated by a multifunctional intron-encoded protein (IEP) that has reverse transcriptase, maturase and DNA endonuclease activities. Homing is the major mobility pathway for group II introns. However, they can be re-targeted to specifically recognize a 14-nt region of DNA juxtaposed to the fixed sites of IEP interaction. This feature can be exploited for the design of new gene therapy strategies. Thus, group II introns could be engineered for insertion into harmful DNA sequences, a technique already used successfully to target extrachromosomal genes in human cells [38]. However, it remains to be seen whether this process works with eukaryotic chromosomal genes. If it does, group II introns might be used to specifically and permanently disrupt genes of interest. Moreover, these introns can also carry foreign genes [209, 210], a capability that might allow them to deliver therapeutic genes to particular sites in the genome. 3.2 Modified ribozymes 3.2.1 Catalytic antisense RNAs The extension of ribozyme substrate recognition arms was the first modification aimed at achieving optimal association between ribozymes and their substrates. This resulted in a new class of ribozymes called catalytic antisense RNAs. These extended arms help ribozymes gain access to target sequences in long RNA substrates. However, the stability of these duplexes prevents the dissociation of the cleavage products, impeding ribozyme turnover (reviewed in [211]). This strategy, mainly used with the hammerhead motif, has been tested for blocking the replication of HIV-1 [212, 213]. The inhibition achieved was 4–7-fold more effective than when using the antisense or ribozyme models alone. Moreover, whereas catalytic antisense RNAs are poorly active in vitro, they are strongly inhibitory ex vivo [213]. These catalytic RNA molecules have also been shown to successfully attenuate the expression of the white gene involved in Drosophila eye pigmentation [214], and of the npt gene in plants [215]. Asymmetric hammerhead ribozymes are made by modifying catalytic antisense RNAs. The modified versions consist of a single long arm that forms helix III. Helix I is reduced to only 3 nt (Fig. 4) [216]. The main advantages of this asymmetric design are the ability to release the 3′-cleavage product (the antisense effect remaining intact) and the easy generation of these catalytic RNAs by PCR [217]. These ideas have been corroborated in in vitro and ex vivo assays in which specific asymmetric hammerhead ribozymes were designed to inactivate RNAs involved in different pathologies, such as Bcr-abl mRNA in chronic myeloid leukemia (CML) [218], interleukin-2 mRNA in autoimmune disorders [219], and tat mRNA of HIV-1 [220]. A novel way of designing catalytic antisense RNAs has recently been reported [221]. In contrast to random extension of the recognition arms, the authors describe the use of a stable stem–loop motif linked to the 3′-end of the ribozyme as an antisense domain. This stem–loop must be complementary to another stem–loop motif (the sense domain) in the target RNA molecule. This design of sense and antisense domains resembles those of naturally occurring antisense-controlled systems in which loop–loop interactions are responsible for the efficient binding of the two RNA molecules [222]. These inhibitor RNAs were developed for both hairpin and hammerhead ribozymes [221, 223]. The efficiency of these newly designed catalytic antisense RNAs has been tested in vitro against HIV-1 TAR-containing RNAs. The antisense stem–loop motif is the complementary sequence of the TAR stem–loop domain. All the catalytic antisense RNAs assayed processed the HIV RNAs more efficiently than the model ribozymes (hairpin or hammerhead), suggesting that the TAR domain might be useful as an anchoring site for HIV ribozyme targeting [223]. These experiments also demonstrated that the two target sequences, the cleavage site and the sense domain, do not need to be contiguous within the substrate RNA. A similar strategy, based on the use of facilitators, pursues the same aim without having to modify ribozymes [224]. This method makes use of external oligonucleotides specifically designed to bind the sequences flanking the target site in the substrate RNA. This renders the cleavage site more accessible to the ribozyme [225]. This strategy has, however, only been tested with hammerhead ribozymes. Facilitator length varies from 12 to 24-nt, and DNA, RNA and modified oligonucleotides have all been reported useful [226, 227]. These facilitators improve hammerhead cleavage and increase the turnover rate [228–231]. However, it is reported that facilitator oligonucleotide inhibits ribozyme activity under certain conditions [232]. To our knowledge, there are no reports on the successful use of this strategy in mammalian cells. 3.2.2 RNPs The interaction or association of RNA molecules with cellular factors (e.g., proteins), is an important difference between in vitro and ex vivo assays. Moreover, essential biological processes are mediated by RNP complexes [233]. These associations are important for achieving RNA stability and RNA catalysis. To date, several proteins have been characterized that bind to hammerhead ribozymes and act as RNA chaperones, promoting the unwinding of the RNA substrate, strand-exchange and annealing, and co-localization of the ribozyme with its specific target. This methodology has been shown to be useful to overcome limitations in specificity and turnover. One of the first strategies developed was based on a protein derived from the p7 nucleocapsid of HIV-1. Retroviral nucleocapsid proteins can bind basic residues of RNA molecules, especially in areas of single-stranded RNA [234]. Further, acting as RNA chaperones, they are thought to be responsible for the destabilization of RNA helices. They can increase the turnover of hammerhead ribozymes by 20–30-fold [235–237] and subsequently increase the cleavage rate. Another protein widely used, and which has given similar results, is A1. This contains three RNA binding domains. Two in the N-terminal region allow binding to single-stranded RNA in a non-specific way, as well as an unwinding activity. The third, which lies in the C-terminal region and is named the RGG domain, determines binding in a cooperative manner as well as the annealing of single-stranded RNA molecules [235]. These features suggest that A1 protein might be involved in in vivo interactions with most cellular RNAs. A1 has been used as an RNA chaperone, improving ribozyme binding specificity and turnover [235, 237]. Another protein shown to bind to a specific hammerhead ribozyme is glyceraldehyde 3-phosphate dehydrogenase (GAPDH) [238, 239]. In contrast to the previous examples, binding to a conformational motif of the ribozyme used in this work occurs. GAPDH accelerates both the formation of the ribozyme–substrate complex and the dissociation of products. This is likely due to an RNA-unfolding activity of GAPDH. A new strategy, exploiting the positive features of RNA binding proteins, has been developed. Chimeric RNA molecules have been designed, which combine a catalytic RNA domain and a decoy domain with the ability to capture proteins that either promote an improvement in the ribozyme performance [240–242] or are essential in viral life cycles [243–245]. Recently, Taira and co-workers have developed a very elegant strategy for overcoming the problem of accessibility of ribozymes to their target sequences in folded RNA substrates [240, 241]. These authors created a chimeric molecule formed of a catalytic domain (hammerhead ribozyme) and a decoy domain (a constitutive transport element [CTE], or a polyA sequence attached to the 3′-end of the ribozyme; Fig. 9). These decoy domains recruit helicase A and eIF4A proteins (which have RNA helicase activity) respectively. The main goal of this strategy is to destabilize complex substrate structures by binding an RNA helicase to the ribozyme. In all tests, the RNA–protein hybrids were more effective as inhibitors than ribozymes without the decoy motif. The reason for this enhancement is probably improved accessibility to targets due to the unwinding activity of the RNA helicase rather than any positive effect on intracellular ribozyme stability. The main advantage of these systems is that they are easy to design and use, making them suitable for in vivo applications. Figure 9 Open in new tabDownload slide Chimeric RNA combining both catalytic and helicase activities. Schematic representation of the mechanism of action of the chimeric RNA molecules including a ribozyme domain and a helicase complex. The inhibitor RNA is represented by a hammerhead ribozyme and the constitutive transport element domain responsible for the binding of the helicase protein complex. The ribozyme-protein hybrid would be able to unwind the local secondary structure of the target RNA, allowing the ribozyme to bind to its specific cleavage site in the substrate. Figure adapted from Warashina et al. [240]. Figure 9 Open in new tabDownload slide Chimeric RNA combining both catalytic and helicase activities. Schematic representation of the mechanism of action of the chimeric RNA molecules including a ribozyme domain and a helicase complex. The inhibitor RNA is represented by a hammerhead ribozyme and the constitutive transport element domain responsible for the binding of the helicase protein complex. The ribozyme-protein hybrid would be able to unwind the local secondary structure of the target RNA, allowing the ribozyme to bind to its specific cleavage site in the substrate. Figure adapted from Warashina et al. [240]. The other strategy reported for combined catalytic and decoy domains is to employ specific binding sequences of viral proteins (as decoy domains) to achieve two inhibitory effects from a single molecule: ribozyme cleavage and the sequestration of a protein essential in viral life cycles. Exploiting the results described above for RNA chaperone proteins, Burke and co-workers developed a revolutionary strategy [111]. They constructed a chimeric hairpin ribozyme, which included a protein-binding domain for the coat protein of bacteriophage R17 as a prolongation of helix 4 (Fig. 5). This protein is involved in the control of gene expression and virus assembly [246]. Far from impeding the activity of the ribozyme, the linkage of this protein-binding domain to the hairpin motif induced a slight increase in activity. Further, the protein remained bound to the ribozyme during catalysis. This preliminary study led other groups to investigate the possibility of using decoy domains to complement the action of ribozymes. One of the most used decoy domains has been the RRE region of HIV. The binding of Rev protein to its specific RRE site is essential for HIV replication. The linkage of a stem–loop aptamer of the RRE motif to a ribozyme induces an in vivo inhibition of virus infectivity [243],[244]. 3.2.3 Allosteric ribozymes Allosteric ribozymes are a modified type of catalytic RNA whose activity can be regulated by external factors. Control of their cleavage activity is achieved by binding an effector molecule to an allosteric binding site (different from the active site). This strategy can be exploited in those situations in which a regulation of RNA-cleavage activity by an external factor is desirable, e.g., cleavage of mRNAs or viral RNAs in infected cells. It has also been used in biotechnology settings, where the allosteric ribozymes, acting as biosensor molecules, detect different analytes. Allosteric ribozymes have been obtained by rational design, in vitro selection, or a combination of both [247–249]. Although hammerhead and hairpin ribozymes have been the catalytic motifs most widely used [247–251], the development of allosteric ribozymes with non-natural catalytic domains has also been reported [248]. Different molecules have been used as effectors, including small molecules, metal ions, proteins and oligonucleotides. Some of the allosteric ribozymes responsive to these molecules are summarized in Table 1. The allosteric mechanisms of different effectors have been widely studied. Nevertheless, the different mechanisms of allosteric regulation include conformational changes modulated by the effector molecule. These changes are the result of steric and antisense interactions, secondary structure stabilizations and quaternary structure modulations [247]. Table 1 Allosteric ribozymes Effector molecule Reference Small molecules ATP [278–280] Flavin mononucleotide [273, 281–283] Theophylin [248, 284] cNMPs [285, 286] Metal ions [276, 287, 288] Proteins [289–291] Oligonucleotides [252, 254, 255, 261, 292] Effector molecule Reference Small molecules ATP [278–280] Flavin mononucleotide [273, 281–283] Theophylin [248, 284] cNMPs [285, 286] Metal ions [276, 287, 288] Proteins [289–291] Oligonucleotides [252, 254, 255, 261, 292] Open in new tab Table 1 Allosteric ribozymes Effector molecule Reference Small molecules ATP [278–280] Flavin mononucleotide [273, 281–283] Theophylin [248, 284] cNMPs [285, 286] Metal ions [276, 287, 288] Proteins [289–291] Oligonucleotides [252, 254, 255, 261, 292] Effector molecule Reference Small molecules ATP [278–280] Flavin mononucleotide [273, 281–283] Theophylin [248, 284] cNMPs [285, 286] Metal ions [276, 287, 288] Proteins [289–291] Oligonucleotides [252, 254, 255, 261, 292] Open in new tab Only oligonucleotide-regulated ribozymes have been used for gene silencing. Different strategies have been developed for oligonucleotide-mediated regulation, but in all cases an inactive form of the ribozyme becomes active in the presence of the oligonucleotide molecule (Fig. 10) [252, 253]. There are reports describing the use of extra sequences within the ribozyme molecule that induce the formation of an inactive conformer. Such extra sequences work as attenuators (Fig. 10a)[250, 252]. The presence of the effector oligonucleotide, complementary to the attenuator, makes the ribozyme adopt the active conformation. Burke and co-workers have developed a system in which the effector molecule does not bind directly to the attenuator sequence (Fig. 10b) [254]. This targeted ribozyme-attenuated probe design requires no prior knowledge of ribozyme structure, and the authors hypothesize that a more generic regulation should be possible if the effector binding sequences and the attenuation sequences are separated into different motifs. Figure 10 Open in new tabDownload slide Oligonucleotide-regulated ribozymes. Schematic representation of different examples of oligonucleotide-regulation of the ribozyme catalytic activity. The oligonucleotide is represented by a thick line. Ribozyme is shown in black and target molecule in gray. a: An extra sequence (attenuator shown in red) within the ribozyme makes inactive the catalytic domain. The oligonucleotide effector specifically binds to the attenuator sequence releasing the catalytic domain and therefore restoring the catalytic performance. b: Schematic representation of the targeted ribozyme-attenuated probe design. The ribozyme activity is sequestered by the binding of an attenuator sequence to the catalytic domain, as described in (a), however the effector oligonucleotide binds to an antisense sequence (shown in blue) different than the attenuator one. As a result of this interaction the catalytic domain gets free of the attenuator and the catalytic activity is restored. c: The effector oligonucleotide binds simultaneously to the ribozyme and the substrate, resulting in the formation of a three-way junction complex that allows ribozyme activity. Figure 10 Open in new tabDownload slide Oligonucleotide-regulated ribozymes. Schematic representation of different examples of oligonucleotide-regulation of the ribozyme catalytic activity. The oligonucleotide is represented by a thick line. Ribozyme is shown in black and target molecule in gray. a: An extra sequence (attenuator shown in red) within the ribozyme makes inactive the catalytic domain. The oligonucleotide effector specifically binds to the attenuator sequence releasing the catalytic domain and therefore restoring the catalytic performance. b: Schematic representation of the targeted ribozyme-attenuated probe design. The ribozyme activity is sequestered by the binding of an attenuator sequence to the catalytic domain, as described in (a), however the effector oligonucleotide binds to an antisense sequence (shown in blue) different than the attenuator one. As a result of this interaction the catalytic domain gets free of the attenuator and the catalytic activity is restored. c: The effector oligonucleotide binds simultaneously to the ribozyme and the substrate, resulting in the formation of a three-way junction complex that allows ribozyme activity. Another way to regulate ribozyme activity is to make substrate-binding dependent on the presence of the RNA or DNA effector. This has been described by Sen and co-workers who indicate that the effector molecule binds simultaneously to both ribozyme and substrate to form a branched, three-way complex (Fig. 10c) [253, 255]. Further, Ohtsuka and co-workers have developed a system in which the activating oligonucleotide stabilizes an otherwise unstable structure [256]. These authors replaced stem II of a hammerhead ribozyme with a large loop of defined sequences, abolishing cleavage activity. The binding of the effector molecule to sequences along one side of the loop allows the stabilization of the active conformation of the ribozyme, making oligonucleotide-controlled activation possible. Finally, a type of allosteric regulation involving an oligonucleotide effector and different aspects of quaternary structure modulation has been developed by Taira and co-workers (reviewed in [257]). Maxizymes are allosteric ribozymes whose activity is regulated by a specific sequence in the target mRNA. This sequence, called the sensor sequence, is physically different from the cleavage site. Maxizymes were first developed as ribozymes to cleave target RNA at two different sites [258]. They are composed of two minizymes (deleted hammerhead ribozymes) which are active only when they form a dimer [259]. They have been successfully used for targeting HIV-1 tat mRNA in mammalian cell culture [260]. Further developments have allowed maxizymes to act as real allosteric ribozymes. The two parts of the molecule have been designed so that one part acts as the sensor molecule and the other as the catalytic molecule. The sensor molecule is often deleted to inactivate the ribozyme, making the molecule even smaller. Only when both parts form the dimer is cleavage activity obtained (Fig. 11). Further, the active dimer is formed only when the two substrate-binding regions properly bind both target sites in the substrate RNA. [257]. Maxizymes targeting different genes have already been constructed and their activity demonstrated [261–263]. A maxizyme against Bcr-abl chimeric mRNA has been widely studied. The sensor targets the junction between the bcr and abl genes while the active molecule targets abl exon 2. The activity of this maxizyme has been shown in ex vivo assays [264], and its antitumorigenic effect in murine CML models has been assessed [263]. Injection of a CML cell line into mice was lethal after 90 days in all cases. However when maxizyme-expressing CML cells were injected, all mice remained disease-free. This is the first report of a successful in vivo application of an allosteric ribozyme. Recently, maxizyme technology has been successfully used to target Bradeion mRNA (specifically expressed in human colorectal cancer and malignant melanoma) both ex vivo and in a mouse system [265]. Figure 11 Open in new tabDownload slide Schematic representation of an active maxizyme. Maxizymes activity is regulated by the presence of a determined sequence (sensor sequence) in the target mRNA. The active conformation is only achieved when both domains (sensor domain and catalytic domain) bind properly to the target RNA. Figure adapted from Kuwabara et al. [257]. Figure 11 Open in new tabDownload slide Schematic representation of an active maxizyme. Maxizymes activity is regulated by the presence of a determined sequence (sensor sequence) in the target mRNA. The active conformation is only achieved when both domains (sensor domain and catalytic domain) bind properly to the target RNA. Figure adapted from Kuwabara et al. [257]. As described for the trans-splicing strategy, maxizymes can specifically target virus-infected cells or be used in a tissue-specific manner. A viral RNA sequence or tissue-specific RNA would be selected as the sensor sequence, and the sequence of an essential cellular gene as the cleavage site. 4 Other ribozyme applications 4.1 Inverse genomics Once the genome projects of organisms have concluded, elucidating the role of the different genes and non-translated sequences, as well as their connections in cellular pathways, becomes the priority. Several strategies have been developed for these investigations [266, 267], but its success has been limited. Ribozymes have been successfully used for determining gene function. Early assays focused on inhibiting specific genes by generating modulatable ‘non-functional’ phenotypes [268], phenotypic variants with reduced levels of mRNAs [269], and complete knock-outs [270, 271]. However, these strategies have a limited ability to clarify the function of genes involved in cellular pathways and for which altered expression patterns lead to complex phenotypes; the use of a ribozyme-based inverse genomic strategy is a better approach. A ribozyme library, in which the substrate recognition sequences have been randomized, is introduced into a cell line (Fig. 12). When any gene involved in the cellular pathway under study is inhibited by the action of an active ribozyme, the cells display a selectable phenotype. Those showing the desired phenotype are rescued and the ribozymes-encoding DNAs isolated and sequenced. The identification of the target is deduced by complementarity to the ribozyme arms, and the resulting sequence used to identify the gene responsible for the phenotype. This strategy has clarified the identity of a wide number of genes (Table 2). However, despite positive results, this technique is susceptible to different factors such as ribozyme expression level, steric hindrance of the ribozyme in its attempts to access the target site, and the cellular compartmentalization of the ribozyme and its substrate, etc. [272]. To overcome the accessibility problem, Taira and co-workers used a chimeric ribozyme-poly(A) tail molecule (see Section 3.2.2) [241]. Figure 12 Open in new tabDownload slide Ribozyme-based inverse genomic approach. Representation of the general mechanism of the inverse genomic approach performed with a randomized ribozyme library. Different methods proposed for the identification of genes are indicated. C-SPACE, is a strategy developed by Wong-Staal and co-workers, specifically designed for the identification of the 5′-products generated by a hairpin ribozyme [304]. Figure 12 Open in new tabDownload slide Ribozyme-based inverse genomic approach. Representation of the general mechanism of the inverse genomic approach performed with a randomized ribozyme library. Different methods proposed for the identification of genes are indicated. C-SPACE, is a strategy developed by Wong-Staal and co-workers, specifically designed for the identification of the 5′-products generated by a hairpin ribozyme [304]. Table 2 Gene function discovery by ribozyme-based inverse genomics Biological processes Gene Reference Cell–cell contacts, cytoskeleton organization Ppan [293] CAP-independent translation mediated by HCV IRES eIF2γ, eIF2Bγ, α-PSMA7 [294, 295] Suppression of cellular transformation mTert [272] Transcription regulation of BRCA1 Id4 [296] Apoptosis tradd, caspase 2, caspase 8, RIP, RAIDD, TRAF2, ETS1, bak [297, 298] Biological processes Gene Reference Cell–cell contacts, cytoskeleton organization Ppan [293] CAP-independent translation mediated by HCV IRES eIF2γ, eIF2Bγ, α-PSMA7 [294, 295] Suppression of cellular transformation mTert [272] Transcription regulation of BRCA1 Id4 [296] Apoptosis tradd, caspase 2, caspase 8, RIP, RAIDD, TRAF2, ETS1, bak [297, 298] Open in new tab Table 2 Gene function discovery by ribozyme-based inverse genomics Biological processes Gene Reference Cell–cell contacts, cytoskeleton organization Ppan [293] CAP-independent translation mediated by HCV IRES eIF2γ, eIF2Bγ, α-PSMA7 [294, 295] Suppression of cellular transformation mTert [272] Transcription regulation of BRCA1 Id4 [296] Apoptosis tradd, caspase 2, caspase 8, RIP, RAIDD, TRAF2, ETS1, bak [297, 298] Biological processes Gene Reference Cell–cell contacts, cytoskeleton organization Ppan [293] CAP-independent translation mediated by HCV IRES eIF2γ, eIF2Bγ, α-PSMA7 [294, 295] Suppression of cellular transformation mTert [272] Transcription regulation of BRCA1 Id4 [296] Apoptosis tradd, caspase 2, caspase 8, RIP, RAIDD, TRAF2, ETS1, bak [297, 298] Open in new tab 4.2 Ribozymes as biosensor molecules Allosteric ribozyme technology is now part of the biotechnology toolbox [273]. The controlled cleavage of a reporter substrate by an allosteric ribozyme might be used for detecting an analyte able to switch on the ribozyme: in other words, ribozymes can be used as biosensor molecules. The desirable characteristics of such biosensor ribozymes have recently been reviewed by Breaker [274]. One of the main advantages of using ribozymes as biosensor molecules is that a single molecule undertakes the two required biosensor activities: molecular recognition and signal generation. Further, ribozymes can be used for both qualitative and quantitative determinations [275]. An array of seven allosteric ribozymes has been used successfully to detect different analytes in a complex mixture [276]. The prototype array was produced by attaching previously characterized self-cleavable allosteric hammerhead ribozymes to a gold surface. Via the detection of cAMP, the same RNA array was used to determine the phenotype of different E. coli strains for adenylate cyclase function [276]. The authors demonstrated that the array could be used for qualitative and quantitative determinations. Famulok and co-workers have described the possibility of using protein-dependent ribozymes to report molecular interactions [251]. They developed hammerhead ribozymes whose activity was modulated (activated or inhibited) by the binding of the HIV Rev protein. These Rev-responsive ribozymes were used to screen a battery of antibiotics via their capacities to bind to Rev and, therefore, to return ribozymes to their initial state. An external substrate labeled with a fluorescent molecule at its 5′-end, and with a fluorescent quencher at its 3′-end, was used as a reporter, allowing real time determinations. The authors identified a molecule able to bind Rev efficiently, which led to the inhibition of HIV-1 replication in cultured cells [251]. Similarly, using either hairpin or hammerhead ribozymes, different protein–RNA, protein–small molecules and protein–protein interactions could be monitored [251]. In addition, Seiwert and co-workers have used allosteric hammerhead ribozymes to detect post-translational modifications of proteins [277]. These authors designed ribozymes whose activity was modulated either by the unphosphorylated form of protein kinase ERK2, or by its phosphorylated form, and were able to quantitatively detect the target protein in complex mixtures such as cell lysates. 5 Concluding remarks The results reported in the literature along the last two decades, which we have tried to summarize in this work, clearly demonstrate the potential applicability of ribozymes as specific gene silencers and their utility as therapeutic and biotechnological tools. Further, many efforts have been dedicated during the last years towards resolving the problems pointed out by the first attempts of using these fascinating molecules inside cells. The innovative reported methods make to conceive great hopes on the potential of these molecules for the development of successful RNA-based therapeutics. Thus, several clinical trials are currently attempting to analyze the efficacy and validity of ribozymes as therapeutic agents. The combination of RNA-based therapies with conventional drugs is hardly encouraged at the moment. The use of in vitro selection techniques to optimize the known ribozymes, to isolate completely new RNA inhibitors, or to develop innovative utilization strategies, will contribute to the establishment of an efficient gene-inactivating technology. Besides the therapeutic uses, other applications can be envisioned. Thus, biotechnology is a completely open field where the ribozyme uses are still almost unexplored. 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TI - Ribozymes: recent advances in the development of RNA tools
JO - FEMS Microbiology Reviews
DO - 10.1016/S0168-6445(03)00020-2
DA - 2003-04-01
UR - https://www.deepdyve.com/lp/oxford-university-press/ribozymes-recent-advances-in-the-development-of-rna-tools-0wY6p71RL1
SP - 75
EP - 97
VL - 27
IS - 1
DP - DeepDyve
ER -