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Mechanism of maturase-promoted group II intron splicing

2001; Springer Nature; Volume: 20; Issue: 24 Linguagem: Inglês

10.1093/emboj/20.24.7259

1460-2075

Manabu Matsuura, James W. Noah, Alan M. Lambowitz,

RNA modifications and cancer

Article17 December 2001free access Mechanism of maturase-promoted group II intron splicing Manabu Matsuura Manabu Matsuura Institute for Cellular and Molecular Biology, Department of Chemistry and Biochemistry, and Section of Molecular Genetics and Microbiology, School of Biological Sciences, University of Texas at Austin, Austin, TX, 78712 USA Search for more papers by this author James W. Noah James W. Noah Institute for Cellular and Molecular Biology, Department of Chemistry and Biochemistry, and Section of Molecular Genetics and Microbiology, School of Biological Sciences, University of Texas at Austin, Austin, TX, 78712 USA Search for more papers by this author Alan M. Lambowitz Corresponding Author Alan M. Lambowitz Institute for Cellular and Molecular Biology, Department of Chemistry and Biochemistry, and Section of Molecular Genetics and Microbiology, School of Biological Sciences, University of Texas at Austin, Austin, TX, 78712 USA Search for more papers by this author Manabu Matsuura Manabu Matsuura Institute for Cellular and Molecular Biology, Department of Chemistry and Biochemistry, and Section of Molecular Genetics and Microbiology, School of Biological Sciences, University of Texas at Austin, Austin, TX, 78712 USA Search for more papers by this author James W. Noah James W. Noah Institute for Cellular and Molecular Biology, Department of Chemistry and Biochemistry, and Section of Molecular Genetics and Microbiology, School of Biological Sciences, University of Texas at Austin, Austin, TX, 78712 USA Search for more papers by this author Alan M. Lambowitz Corresponding Author Alan M. Lambowitz Institute for Cellular and Molecular Biology, Department of Chemistry and Biochemistry, and Section of Molecular Genetics and Microbiology, School of Biological Sciences, University of Texas at Austin, Austin, TX, 78712 USA Search for more papers by this author Author Information Manabu Matsuura1, James W. Noah1 and Alan M. Lambowitz 1 1Institute for Cellular and Molecular Biology, Department of Chemistry and Biochemistry, and Section of Molecular Genetics and Microbiology, School of Biological Sciences, University of Texas at Austin, Austin, TX, 78712 USA ‡M.Matsuura and J.W.Noah contributed equally to this work *Corresponding author. E-mail: [email protected] The EMBO Journal (2001)20:7259-7270https://doi.org/10.1093/emboj/20.24.7259 PDFDownload PDF of article text and main figures. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Mobile group II introns encode reverse transcriptases that also function as intron-specific splicing factors (maturases). We showed previously that the reverse transcriptase/maturase encoded by the Lactococcus lactis Ll.LtrB intron has a high affinity binding site at the beginning of its own coding region in an idiosyncratic structure, DIVa. Here, we identify potential secondary binding sites in conserved regions of the catalytic core and show via chemical modification experiments that binding of the maturase induces the formation of key tertiary interactions required for RNA splicing. The interaction with conserved as well as idiosyncratic regions explains how maturases in some organisms could evolve into general group II intron splicing factors, potentially mirroring a key step in the evolution of spliceosomal introns. Introduction Group II introns found in organelle and bacterial genomes splice via a lariat intermediate and are thought to be ancestors of nuclear pre-mRNA introns (Michel and Ferat, 1995). The group II intron splicing reactions are RNA catalyzed and require a conserved RNA structure formed by six interacting double-helical domains (DI–VI; Michel and Ferat, 1995). An evolutionary relationship between group II and spliceosomal introns is supported by similarities in reaction mechanisms and splice site consensus sequences, by analogies between the structure and function of group II intron domains and snRNAs, by the ability of the 'snRNA-like' group II intron domains to associate in trans and by the discovery that some group II introns are mobile genetic elements, suggesting how introns could have spread in nuclear genomes (Michel and Ferat, 1995; Lambowitz et al., 1999). Together, these findings suggest a scenario in which mobile group II introns invaded nuclear genomes and then degenerated, with group II intron domains evolving into snRNAs. If so, further studies of mobile group II introns may provide insight into how present-day splicing mechanisms evolved. The best studied mobile group II introns are the yeast mtDNA introns aI1 and aI2 and the Lactococcus lactis Ll.LtrB intron (reviewed in Lambowitz et al., 1999). These introns encode reverse transcriptases that function both in intron mobility and as maturases in RNA splicing. The group II intron-encoded proteins (IEPs) are typically encoded in DIV, outside the intron's catalytic core, and have three major conserved domains: an N-terminal reverse transcriptase domain, domain X associated with splicing or maturase activity and a C-terminal DNA endonuclease domain, which functions in intron mobility. After translation, the IEP binds specifically to the intron RNA to promote splicing and then remains associated with the excised intron lariat RNA to form a ribonucleoprotein (RNP) complex that catalyzes intron mobility. In the major mobility pathway, retrohoming, the intron RNA in this RNP reverse splices directly into a specific target site in double-stranded DNA and is reverse transcribed by the IEP. In addition to homing, group II introns can also retrotranspose to ectopic sites that resemble the normal homing site at low frequency, leading to the dispersal of the introns to different genomic locations (Dickson et al., 2001; and references therein). The splicing activity of the group II IEPs was first demonstrated genetically for the two yeast mtDNA introns and subsequently both genetically and biochemically for the L.lactis Ll.LtrB intron (see Lambowitz et al., 1999). All three IEPs function specifically in splicing the intron in which they are encoded, although the yeast aI1 and aI2 maturases appear to have some cross-reactivity. Other group II introns encode related proteins that have, in some cases, been inferred to function in splicing. Many of these proteins contain all three conserved domains, but some lack the C-terminal endonuclease domain, while others, such as the MatK proteins encoded by chloroplast tRNALys introns, retain only a well conserved domain X and remnants of the reverse transcriptase domain (Mohr et al., 1993). The latter proteins are presumed to have lost dispensable mobility functions, but retain the essential splicing function. Although all three characterized maturases are intron-specific splicing factors, several investigators have suggested that putative maturases in some organisms function in splicing multiple group II introns. Suggestive evidence for such a generalized role of group II intron maturases comes from experiments showing that inhibition of chloroplast protein synthesis in higher plants blocks the splicing of multiple group II introns, including some that do not themselves encode maturases (Vogel et al., 1999; and references therein), and that maturase-like proteins, no longer encoded within introns, are conserved in the chloroplast DNAs of non-photosynthetic plants that still retain introns in essential genes (Wolfe et al., 1992; Copertino et al., 1994). Highly degenerate Euglena group II introns, referred to as group III introns, are also hypothesized to use a common splicing apparatus that includes a maturase encoded by one of the introns (Copertino et al., 1994). The evolution of an intron-specific maturase to function in splicing multiple group II introns may parallel a step in the evolution of a common cellular splicing machinery for spliceosomal introns. To study the maturase-promoted splicing reaction biochemically, we have developed an experimental system based on the L.lactis Ll.LtrB intron, where it is possible to obtain large amounts of the purified IEP (denoted LtrA protein) via expression in Escherichia coli (Matsuura et al., 1997). We showed that this purified maturase is sufficient to promote the splicing of the Ll.LtrB intron in vitro at low Mg2+ concentrations, where the intron RNA cannot by itself fold into the catalytically active RNA structure (Saldanha et al., 1999). The protein binds specifically to the Ll.LtrB intron, but not to other group II introns, as expected for an intron-specific splicing factor. Deletion analysis showed that the LtrA protein has a primary high affinity binding site in intron subdomain DIVa, an idiosyncratic structure near the beginning of its own coding region, with other regions of DIV also contributing to binding (Wank et al., 1999). Although LtrA does not bind tightly to the intron when DIV is deleted, its binding to DIV is enhanced by both upstream and downstream regions of the intron, and the interaction of LtrA with DIV positions the reverse transcriptase active site to initiate cDNA synthesis just downstream of the intron in the 3′ exon, as required for intron mobility. These findings suggested that after binding to DIV, the maturase might make weaker secondary contacts with 5′ and 3′ regions of the intron to promote the folding of the catalytic core. Supporting this hypothesis, intron RNAs deleted for the primary high affinity binding site in DIVa show residual maturase-dependent splicing in vitro and in vivo, indicating that the weaker, secondary contacts are sufficient to promote splicing even in the absence of high affinity binding to domain IV (Wank et al., 1999; X.Cui and A.M.Lambowitz, in preparation). Here, we analyzed the interaction of the maturase with the Ll.LtrB RNA in detail by quantitative binding assays and RNA footprinting. Our results identify potential secondary binding sites in conserved regions of the catalytic core and demonstrate directly that the binding of the maturase induces an RNA conformational change that results in the formation of key tertiary interactions required for catalytic activity. These findings show how the maturase functions and have implications for the evolution of splicing mechanisms. Results Specific binding of the maturase to 5′ and 3′ segments of the group II intron Figure 1A shows a secondary structure model for the 902 nucleotide ΔORF derivative of the Ll.LtrB intron used in this study. The DIV subsegment of this intron, shown in detail in Figure 1A, contains the high affinity maturase-binding site in DIVa. An in vitro transcript containing the ΔORF Ll.LtrB intron and flanking exons is referred to as Ll.LtrB RNA (see Materials and methods). To detect weaker, secondary binding sites, we first carried out equilibrium binding assays with a derivative of the Ll.LtrB RNA that has a minimal DIV deleted for both DIVa and DIVb (ΔDIVa/b; Figure 1B). In these experiments, small amounts of 32P-labeled RNAs were incubated with increasing concentrations of LtrA protein, and binding was assayed by retention of the complex on a nitrocellulose filter. The assays were done under previously established protein-dependent splicing conditions in reaction medium containing 450 mM NaCl and 5 mM Mg2+ at 30°C, the relatively high salt concentration being necessary to minimize non-specific binding of the LtrA protein (Saldanha et al., 1999). Figure 1.Secondary structure model of the Ll.LtrB intron and equilibrium binding assays. (A) Predicted secondary structure of the 902 nucleotide ΔORF derivative of the Ll.LtrB intron. The structure is based on the generalized group IIA intron structure in Toor et al. (2001). Nucleotide residues involved in the EBS–IBS pairings and tertiary interactions (Greek letters) are marked. Tertiary interactions are modeled based on sequence and structure comparisons with other group II introns and have not been verified experimentally for the Ll.LtrB intron. The η–η′ interaction does not fit the consensus for other group IIA introns and was best modeled as a 'group IIB-type' interaction (Chanfreau and Jacquier, 1996; Costa et al., 1997). The ζ receptor for the unusual GAGA tetraloop at the top of DV (ζ′) is shown as an 11 nucleotide sequence based on the 11 nucleotide GAAA receptor (Costa and Michel, 1995). The η, η′ and ζ elements are indicated in dashed lines to denote these uncertainties. The boundaries of deletions or subsegments used in different experiments are demarcated by brackets, and the branch point adenosine deleted in the ΔA2486 mutant is boxed. The predicted secondary structure of DIV is shown above, with the location of the deleted 1596 nucleotide LtrA ORF segment indicated to the upper right. Subdomain DIVa, the high affinity binding site for the LtrA protein, contains the putative Shine–Dalgarno (SD) sequence (underlined) and initiation codon (boxed) of the LtrA ORF. (B and C) Equilibrium binding assays. 32P-labeled RNAs were incubated with increasing concentrations of LtrA protein for 1 h at 30°C in reaction medium containing (B) 450 mM NaCl/5 mM MgCl2 or (C) 500 mM NH4Cl/5 mM MgCl2. Binding was assayed by the retention of the 32P-labeled RNA on a nitrocellulose filter. The Kd values are the mean ± SD for three experiments for each RNA. The specific binding of LtrA to the ΔDIVa/b intron was not detected previously in reaction media containing bovine serum albumin (Wank et al., 1999). *The Kd value for the wild-type Ll.LtrB RNA is too low to be determined by equilibrium binding; the value 0.25 pM indicated in the figure is the apparent Kd measured as koff/kon (Wank et al., 1999). Download figure Download PowerPoint Figure 1B shows that LtrA does in fact bind specifically to the ΔDIVa/b RNA with an apparent Kd of 18 ± 4 nM. This binding was much weaker than that of the wild-type Ll.LtrB RNA, containing the high affinity binding site (apparent Kd measured as koff/kon = 0.25 pM; Wank et al., 1999), but significantly better than that of a non-specific RNA containing the Neurospora crassa mt LSU group I intron. A 5′ segment of Ll.LtrB RNA consisting of E1-DI also bound specifically with an apparent Kd of 45 ± 8 nM, while a 3′ segment consisting of DV-E2 bound less tightly (apparent Kd >70 nM), but again more strongly than the non-specific RNA. In contrast, a segment containing only DII and DIII did not bind better than the non-specific RNA. Together, these findings show that LtrA can bind specifically to the Ll.LtrB RNA in the absence of its high affinity DIVa binding site and suggest at least two secondary binding sites in the 5′ and 3′ segments of the Ll.LtrB RNA. RNA footprinting strategy To investigate the interaction of the LtrA protein with the intron RNA in detail, we carried out RNA footprinting, using iodine cleavage of phosphorothioate-containing RNAs to assess phosphodiester backbone interactions, and chemical probing with dimethyl sulfate (DMS) and kethoxal to assess base interactions. In general, the binding of a protein can result in RNA protection directly, or indirectly by inducing the formation of higher order RNA structure. To help distinguish these possibilities, we compared modification protection patterns of the Ll.LtrB RNA under protein-dependent and self-splicing conditions in the presence or absence of the LtrA protein. Protections observed in the absence of LtrA at low or high Mg2+ concentration are assigned to RNA structure. Potential protein protection sites were identified as positions that remained accessible in the folded RNA under self-splicing conditions and were protected only in the presence of the LtrA protein, with the best candidates assigned based on additional structural information (see Discussion). The reaction conditions were modified slightly, with NH4+ instead of Na+ as the monovalent cation to optimize self-splicing. Figure 2 shows splicing time courses for the wild-type Ll.LtrB intron under the different conditions used in RNA footprinting. Under protein-dependent conditions (0.5 M NH4Cl/5 mM MgCl2), the wild-type Ll.LtrB RNA showed no detectable splicing in the absence of LtrA, but was spliced efficiently in its presence (Figure 2A and B). The protein-dependent splicing reaction was biphasic, with the fast phase (69%) occurring at 3.3/min and the slow phase (29%) at 0.01/min. The biphasic kinetics under protein excess conditions were attributed to a more slowly folding RNA conformer (Saldanha et al., 1999). When the Mg2+ concentration was increased (0.5 M NH4Cl/50 mM MgCl2), the intron RNA folds into the active structure and self-splices in the absence of LtrA (Figure 2C). This self-splicing occurred at a rate of 0.02/min, considerably slower than protein-dependent splicing at 5 mM Mg2+. LtrA could still bind under these conditions (koff = 0.001/min compared with 0.002/min at 5 mM Mg2+) and increased the reaction rate ∼25-fold [fast phase, 0.5/min (67%); slow phase, 0.06/min (23%); Figure 2D]. Finally, under higher salt conditions (1.5 M NH4Cl/50 mM MgCl2), the rate of self-splicing was maximal (0.025/min; Figure 2E), but the LtrA protein had no effect, presumably reflecting that its binding is impeded under these conditions (not shown). We note that even under the higher salt conditions, the maximal rate of self-splicing is still ∼130-fold slower than the rate of protein-dependent splicing at 5 mM Mg2+. Figure 2.RNA splicing of the Ll.LtrB intron under protein-dependent and self-splicing conditions. Splicing time courses were carried out with 32P-labeled Ll.LtrB RNA (20 nM) at 30°C in different reaction media used for RNA structure mapping and footprinting. (A and B) 0.5 M NH4Cl/5 mM Mg2+ (protein-dependent conditions) in the absence or presence of 200 nM LtrA, respectively. (C and D) 0.5 M NH4Cl/50 mM Mg2+ (self-splicing conditions) in the absence or presence of 200 nM LtrA, respectively. (E) 1.5 M NH4Cl/50 mM Mg2+ (maximal self-splicing conditions). Download figure Download PowerPoint To prevent splicing during footprinting, we used a mutant Ll.LtrB RNA deleted for the branch point adenosine in DVI (ΔA2486). This mutation inhibits splicing completely, with the mutant RNA showing only slow residual hydrolysis at the splice sites ( 25% decrease in band intensity). Large and small blue or open triangles indicate positions strongly (>50%) or moderately (49–25%) protected by RNA structure at 50 mM Mg2+ (lanes 4 and 5) compared with 5 mM Mg2+ (lane 1). Blue triangles indicate positions protected at 50 mM Mg2+ (lanes 3–5) and also protected at 5 mM Mg2+ in the presence of LtrA (lane 2), presumably reflecting the formation of all or part the active RNA structure in the presence of the protein. Open triangles indicate a subset of positions protected at 50 mM Mg2+ (lanes 3–5), but not at 5 mM Mg2+ in the presence or absence of LtrA (lanes 1 and 2), possibly reflecting the formation of aberrant RNA structures or interactions at 50 mM Mg2+. Large and small red triangles (protein-dependent protections) indicate positions that remained accessible at 50 mM Mg2+ (lanes 3 and 5) and were strongly (>50%) or moderately (49–25%) protected only in the presence of LtrA protein (lanes 2 and 4). + indicates positions that were protected (>25%) in the free RNA at 50 mM Mg2+ (lane 3) and showed increased protection after addition of LtrA (lane 4), reflecting either strengthened RNA structure or additional protection by the protein. Assignments are based on quantiation of two complete data sets for all positions and three data sets for the DV–VI region. All positions refer to the 5′ phosphate of the marked nucleotide residue. Positions that could not be monitored because of consistent strong reverse transcription stops were C-6, C-1, G1, A393, U2322, U2328, U2340, U2365 and U2375. Download figure Download PowerPoint First, the results show that some regions of phosphodiester backbone are protected by RNA structure, even under protein-dependent conditions (0.5 M NH4Cl/5 mM Mg2+). These regions, highlighted in gray shading in Figure 3B, are found primarily in DI and DII, with additional protections scattered in DIII, DIV and DVI. Under self-splicing conditions (0.5 M NH4Cl/50 mM Mg2+), many of these protections were intensified and a number of additional positions in DI, DII, DIII, DV and DVI were protected (Figure 3, blue or open triangles). The protection pattern was essentially the same under the high salt conditions (1.5 M NH4Cl/50 mM Mg2+). In general, the regions that were protected at 50 mM Mg2+ (blue or open triangles plus gray shading) neighbor sites of known tertiary interactions and include DIb (α), DIc1 (θ and ϵ′), DIc2 (β), DId(i)–(iii) (ζ and κ), DId2 (β′), DId3 (α′), DII (η and θ′), one side of DIII, parts of DV (κ′, λ′ and ζ′) and the top of DVI (η′). Together, these results suggest that some tertiary structure forms at low Mg2+ concentrations, but that higher Mg2+ concentrations are required to support additional tertiary structure, including many of the long-range interactions between different domains. Most positions that were protected in the free RNA at 50 mM Mg2+ were protected to a similar extent at 5 mM Mg2+ in the presence of LtrA (Figure 3, blue triangles), presumably reflecting the formation of all or part of the active RNA structure in the presence of the protein. However, a few positions (open triangles), mostly in domain V, were protected at 50 mM Mg2+ (Figure 3A, lanes 3–5), but not at 5 mM Mg2+, in either the presence or absence of LtrA (lanes 1 and 2). These protections may reflect the formation of aberrant phosphate backbone structures or interactions at 50 mM Mg2+ that could contribute to the slow rate of self-splicing compared with protein-dependent splicing. We also note that DV is only partially protected at 5 mM Mg2+ in the presence of LtrA, even though most other tertiary structure has formed. In constrast, in a ribozyme derived from the yeast aI5γ intron, DV was completely protected from hydroxyl radical cleavage in reaction medium containing 500 mM KCl/100 mM Mg2+ (Swisher et al., 2001). This discrepancy could reflect differences between group IIA and IIB introns or the nature of the RNA constructs, a different stage of the reaction trapped by the branch point deletion in our construct and/or the formation of aberrant or inappropriately rigid RNA structures at high Mg2+. Superimposed over the RNA protections, candidate protein protection sites were identified as positions that remained accessible at 50 mM Mg2+ and were protected only in the presence of LtrA protein (red triangles). These sites were found mainly in DI, DIV and DVI, with a few additional sites in DII. In DIV, most of the putative protein protection sites were clustered in the previously identified high affinity binding site DIVa, with some additional sites scattered in the DIV stem, L4, DIVb, DIVb1 and DIVb2, in agreement with the finding that these regions also contribute to LtrA binding (Wank et al., 1999). In DI, sites protected in the presence of LtrA protein were found in subdomains b, c1, c2, d(ii)–(iii) and d4, with a few additional sites scattered elsewhere. DIb, c1, c2 and d(ii)–(iii) are conserved group II intron structures involved in long-range tertiary interactions (α–α′, β–β′, ϵ–ϵ′, θ–θ′ and κ–κ′), while DId4 is an idiosyncratic structure found only in subgroup IIA introns (Toor et al., 2001). Outside of DI, other clusters of protein-induced protections were found in DIIb and in DVI on the side opposite the branch point adenosine. Somewhat surprisingly, no phosphate backbone protections were observed in proximal regions of the 5′ or 3′ exons. As indicated previously, binding of the protein positions the reverse transcriptase active site to initiate reverse transcription in the 3′ exon (Wank et al., 1999), but this interaction may require the presence of an appropriate DNA primer. Chemical modification with DMS and kethoxal Figure 4 shows chemical modification experiments with DMS and kethoxal under the same conditions, with modification sites mapped as reverse transcription stops one base prior to the modified nucleotide. Reverse transcription detects DMS modification at the N1 position of unpaired adenines and the N3 position of unpaired cytidines, and kethoxal modifications at the N1 and N2 positions of unpaired guanines. Almost all of the DMS modifications were at adenines, due to inefficient modification of cytidines under the conditions used. Figure 4.DMS and kethoxal modification in the presence or absence of LtrA protein. Ll.LtrB-ΔA2486 RNA (20 nM) was incubated in different reaction media at 30°C in the presence or absence of 50 nM LtrA protein, and then modified with DMS or kethoxal. Modification sites were mapped by primer extension, using 5′-labeled primers complementary to different positions in Ll.LtrB RNA. (A) Representative gels. Lanes: A, dideoxy A sequencing ladder obtained using the same 5′-labeled primers; '–DMS' or '–kethoxal', RNA incubated without DMS or kethoxal; Denatured, RNA modified under denaturing conditions (see Materials and methods). Lanes 1–5, modification reactions; lanes 1 and 2, 0.5 M NH4Cl/5 mM Mg2+ (protein-dependent conditions) in the absence and presence of LtrA, respectively; lanes 3 and 4, 0.5 M NH4Cl/50 mM Mg2+ (self-splicing condition) in the absence and presence of LtrA, respectively; lane 5, 1.5 M NH4Cl/50 mM Mg2+ (maximal self-splicing conditions). Landmarks in the intron are indicated to the right of the gel. (B) Summary of DMS and kethoxal modification protection patterns. Long and short arrows indicate strong and moderate modifications at 5 mM Mg2+ (>10-fold and 2- to 10-fold increase in band intensity compared with the unmodified lane, respectively). Large and small blue circles indicate positions strongly (>50%) or moderately (49–25%) prot