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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, Texas 78712, USA
Reprint requests to: 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; e-mail: lambowitz{at}mail.utexas.edu; fax: (512) 232-3420.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Keywords: ribozyme; RNA–protein interaction; RNA structure; translational regulation
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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; Belfort et al. 2002; Lambowitz and Zimmerly 2004). The best characterized mobile group II introns are the yeast mtDNA introns aI1 and aI2 and the Lactococcus lactis Ll.LtrB intron. Studies with these introns showed that the intron-encoded protein (IEP) binds specifically to the intron in unspliced precursor RNA to promote RNA splicing, and then remains bound to the excised intron lariat RNA in a ribonucleoprotein (RNP) particle that mediates intron mobility (Zimmerly et al. 1995a,b; Yang et al. 1996; Matsuura et al. 1997; Saldanha et al. 1999). RNPs initiate mobility by binding to DNA and recognizing a 30–35-bp DNA target site, with both the IEP and base pairing of the intron RNA contributing to DNA target site recognition. The intron RNA then reverse splices directly into one strand of the duplex DNA, while the IEP cleaves the opposite strand and uses the cleaved 3' end as a primer for reverse transcription of the reverse spliced intron RNA. Genomic integration of the resulting cDNA occurs via host cell DNA recombination or repair pathways, which can differ in different hosts (Eskes et al. 1997, 2000; Cousineau et al. 1998).
The binding of the IEP to the intron RNA is critical for both RNA splicing and intron mobility. Previous studies with the L. lactis Ll.LtrB intron suggested a model in which its IEP, denoted LtrA protein, binds first to a high-affinity binding site in intron subdomain DIVa, an idiosyncratic structure that lies outside the catalytic core and contains the ribosome binding site and initiation codon of the LtrA open reading frame (ORF; Wank et al. 1999). Experiments using reporter gene constructs showed that the binding of LtrA to DIVa down-regulates translation, thereby preventing the accumulation of excess IEP and halting ribosome entry into the intron, which might otherwise impede RNA splicing (Singh et al. 2002). After binding to DIVa, LtrA makes weaker secondary contacts with conserved catalytic core regions to stabilize the active structure for RNA splicing and then remains associated with the excised intron RNA, presumably using some or all of the same interactions, to constitute the RNPs that function in intron mobility (Wank et al. 1999; Matsuura et al. 2001). RNA footprinting and structure mapping experiments identified potential secondary binding sites in DI, DII, and DVI and demonstrated that binding of LtrA stabilizes conserved tertiary structure required for catalytic activity (Matsuura et al. 2001).
Although DIVa is required for maximal RNA splicing, an Ll.LtrB intron deleted for DIVa still shows some residual maturase-dependent RNA splicing, which occurs by the direct binding of the IEP to the catalytic core (Wank et al. 1999; Matsuura et al. 2001; Cui et al. 2004). DIVa contributes to RNA splicing by increasing the binding affinity for the maturase by ~105, and the binding of the maturase to DIVa also appears to facilitate RNA structural transitions that are rate limiting for splicing at saturating protein concentrations (Wank et al. 1999; Matsuura et al. 2001). Additionally, binding to DIVa helps position the IEP to initiate cDNA synthesis just downstream from the intron in the 3' exon, as required for intron mobility (Wank et al. 1999).
The yeast aI2 maturase also has a high-affinity binding site in DIVa, which is required for the stable binding of its IEP in vivo (Huang et al. 2003). This finding is significant because the yeast aI2 maturase is translated as a pre-protein with the upstream exons, and DIVa is no longer used for translation initiation, thus separating its role in that process from its other critical functions. The deletion of DIVa from the yeast aI2 intron inhibited RNA splicing, but the level of residual splicing in vivo in the absence of DIVa was considerably higher than that for the Ll.LtrB intron (70% and 6%–10%, respectively; Huang et al. 2003; Cui et al. 2004). In both the yeast and lactococcal introns, however, the deletion of DIVa almost completely abolished intron mobility, suggesting that the DIVa contacts are more critical for that process than they are for RNA splicing (D’Souza and Zhong 2002; Huang et al. 2003).
The structure of the DIVa binding site in the Ll.LtrB intron was investigated previously by in vitro selection and mutagenesis (Singh et al. 2002). These experiments used a small (119-nt) RNA consisting of DIVa plus the DIV stem loop (denoted DIV+IVa RNA), with nucleotide residues 537–598 "doped" 30% (i.e., synthesized with 10% of each non-wild-type nucleotide residue at each position; Singh et al. 2002). The results showed that the LtrA binding site consists of a small terminal stem–loop (iii) emanating from a purine-rich internal loop, with both sequence and secondary structure contributing to LtrA binding (Fig. 1A). Seven nucleotide residues, shown in red in Figure 1A, were found to be invariant in tightly binding DIVa clones, including five in stem–loop (iii) and A556 and G557 in the purine-rich internal loop. The latter nucleotide residues along with those in stem (iii) are part of the predicted Shine–Dalgarno (SD) sequence for the LtrA ORF. Nucleotide substitutions for each of the invariant nucleotide residues inhibited LtrA binding, with the strongest effects found for mutations in A563 and A564 in loop (iii). The structure of the binding site suggests that LtrA binding down-regulates translation by occluding the Shine–Dalgarno sequence, either directly or by stabilizing RNA secondary structure or both (Singh et al. 2002).
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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shows the predicted secondary structure of DIV+IVa RNA based on the previous in vitro selection in which positions 537–598 were "doped" 30% (Singh et al. 2002). This selection established the structure of the terminal stem–loop (iii) and identified the seven invariant nucleotide residues shown in red, but left a large purine-rich internal loop (iii)/(ii), for which no secondary structure could be predicted. To investigate the structure of this internal loop, we carried out iterative in vitro selection with a pool of 136-nt DIV+IVa RNAs in which the terminal stem–loop and the seven invariant nucleotide residues from the original selection were fixed, whereas the remaining 17 nt in the purine-rich loop were randomized (indicated in green).
The progress of the selection is summarized in Table 1. In the first round, 200 nM of the initial DIVa RNA pool (pool 0) containing 6 x 1012 variants was incubated with 150 nM of purified LtrA protein. The bound RNAs were then collected by nitrocellulose filter binding and reamplified via RT-PCR for the next round of selection. In subsequent rounds, 200 nM RNA pool and 10 nM LtrA protein were used for the binding reaction. The proportion of bound RNAs, monitored by 32P-labeled RNA retained on the filter, increased from 0.25% for pool 1 to 0.68%–0.75% for pools 6 and 7, compared to 0.50% for the wild-type RNA under the same conditions. The low proportion of bound wild-type RNA is thought to reflect that the small RNA easily misfolds into an inactive structure (Wank et al. 1999).
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summarizes the sequences and binding strengths of 46 variants cloned after seven rounds of selection. Each variant has a unique sequence for the 17 randomized positions, with no insertions or deletions. Their binding ratios relative to the wild-type DIV+IVa RNA ranged from 7.5 to 0.2, with all but three of the variants binding as well as or better than the wild-type RNA.
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, respectively. This secondary structure contains an extended stem (ii) with the invariant A556 bulged out of the stem, leaving a smaller bulged loop (iii)/(ii) on the opposite side. In most sequences, the alignment to maintain the G557:C base pair required reducing the size of the bulged loop (iii)/(ii) relative to the wild-type structure. In the wild-type structure, A553 is also bulged, but this feature was not conserved in most clones and in fact appears to be selected against in tightly binding variants (see below).
The predicted secondary structure of the top of stem (ii) is supported by the selection for base pairing at each position (Table 2). Thus, positions 558:580 above G557:C581 could form a Watson–Crick or wobble UG base pair in all of the variants. Further, the base pair at 558:580 is GC or CG in 40 of 46 variants, and the putative partner of the invariant G557 is always C rather than U. As a result, most of the selected variants have two strong base pairs above the bulged A556. Below A556, positions 555:582 are also base paired in all the variants, whereas positions 554:583 are paired in 38 of 46 variants, with seven of the eight at which this position is unpaired compensating by having a GC or CG pair at positions 555:582. Additionally, there was strong selection against a second A residue at position 555 just below A556, and against a U residue between positions 580 and 582. The latter avoids placing a potential base-pairing partner opposite the bulged A556 and also favors strong GC base pairs above A556.
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; Gutell et al. 1992). Finally, although A553 is bulged in the wild-type sequence, there was strong selection against this feature with only two of the more weakly binding variants (40 and 46) retaining this bulged A. Instead, A553 is replaced by another nucleotide residue in 42 of 46 variants, and the nucleotide residue at position 553 is base paired in 37 of 46 variants, including all of the most tightly binding (1–12). The selection against the bulged A553 present in the wild-type structure could reflect that it decreases the stability of the stem and/or interferes with recognition of the bulged A556, which may be critical for LtrA binding.
Effect of mutations in the bulged A residues on LtrA binding
To test the importance of the two bulged A residues for LtrA binding, we made mutant DIV+IVa constructs in which each of these A residues was either deleted or paired with a U residue inserted on the opposite side of the stem. The mutant DIV+IVa RNAs were then tested for their ability to bind LtrA by a competition binding assay with an electrophoretically distinguishable wild-type RNA. As shown in Figure 3, mutations that deleted or paired the bulged A556 (
A556 and A556:U, respectively) strongly decreased the binding affinity for LtrA (binding ratios 0.05 and 0.04, respectively). By contrast, mutations that deleted or paired the bulged A553 (
553 and A553:U) increased the binding affinity for LtrA (binding ratios 2.5 and 2.8, respectively), as expected from the selection against this feature. These findings demonstrate that the bulged A556 is important for LtrA binding and that the bulged A553 antagonizes LtrA binding.
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ORF derivative of the Ll.LtrB-intron is linked to the expression of GFP (Fig. 4A; Cui et al. 2004). The ltrB/GFP fusion is cloned downstream from a phage T7lac promoter, and the LtrA protein preceded by a Shine–Dalgarno sequence is expressed from a position just downstream from the GFP coding sequence. This arrangement makes it possible to introduce mutations into DIVa of the Ll.LtrB-ORF intron without affecting the expression of LtrA. Splicing was assayed in Escherichia coli HMS174(DE3), which contains an integrated isopropyl-thio-ß-D-galactoside (IPTG)-inducible T7 RNA polymerase, by measuring single-cell fluorescence using a fluorescence-activated cell sorter (FACS). The FACS assay, which measures production of spliced product, was shown previously to give results in close agreement with both RT-PCR and poison primer extension assays of in vivo splicing (Cui et al. 2004).
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, IPTG induction of cells containing the wild-type construct resulted in an almost 10-fold increase in GFP fluorescence indicative of in vivo splicing (83 ± 15 units compared to 8.7 ± 0.6 units with and without IPTG induction, respectively). The deletion or pairing of A556, which results in weaker binding of LtrA, appeared to somewhat increase the level of in vivo splicing (140 ± 15 and 110 ± 41 units, respectively), whereas the deletion or pairing of A553, which results in stronger binding of LtrA, appeared to slightly decrease the level of in vivo splicing (68 ± 13 and 77 ± 9 units, respectively). Although these effects are relatively small, the same trends were observed for mutations in both bulged A residues in poison primer extension assays of in vivo splicing (Fig. 5). By comparison, the mutations A563C and A564C in the two invariant A residues in loop (iii), which were shown previously to inhibit LtrA binding (Singh et al. 2002), more strongly inhibited splicing (23 ± 6 and 11 ± 1 units, respectively; Fig. 4), and the complete deletion of DIVa was shown previously to inhibit in vivo splicing in this assay by 90%–94% (Cui et al. 2004). The larger effects of the loop (iii) mutations could reflect that they result in a greater decrease in the binding affinity for LtrA than do the bulged A mutations. The larger effects could also reflect that the loop nucleotide residues are more critical than those in the stem for positioning LtrA to interact with the catalytic core, or that the loop and deletion mutations have a greater effect on the underlying ribozyme activity of the intron RNA.
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ORF intron with a phage T7 promoter inserted near its 3' end is expressed from a donor plasmid and integrates into a recipient plasmid target site cloned upstream of a promoterless tetR gene, thereby activating that gene (Guo et al. 2000). The intron donor plasmid pACD2 used in these experiments expresses the LtrA ORF from a position downstream from the 3' exon, again enabling us to introduce mutations into the DIVa structure within the intron without affecting the expression of LtrA (Karberg et al. 2001).
As summarized in Table 3, the deletion or pairing of the bulged A556, which results in weaker binding of LtrA, significantly inhibited intron mobility (mobility frequencies 27 ± 3.5 and 9.3 ± 2.6% wild type for A556 and A556:U, respectively). Surprisingly, however, the deletion or pairing of the bulged A553, which results in tighter binding to DIVa RNAs, also inhibited intron mobility (mobility frequencies 25 ± 3.7 and 50 ± 14% wild type for A553 and A553:U, respectively), as did the mutations A563C and A564C in loop (iii) (mobility frequencies 26 ± 9.8 and 1.5 ± 1.4% wild type, respectively). By comparison, deletion of the entire DIVa stem–loop reduced the mobility frequency to 6 x 10–6 in this assay (D’Souza and Zhong 2002).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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ORF intron (Matsuura et al. 2001). These experiments showed that the bulged A556, as well as all A and G residues in the two base pairs above and below A556 (G557, G588, A582, and A583) are accessible to dimethylsulfate or kethoxal modification in the free RNA at 5 or 50 mM Mg2+, but strongly protected under these conditions by LtrA binding (Matsuura et al. 2001). Additionally, the protections by LtrA extend throughout stem (ii), with the bulged A553 being moderately protected. By contrast, A and G residues in the predicted purine-rich loop (A570–A579) were accessible in the free RNA at 5 mM Mg2+, moderately protected by RNA structure at 50 mM Mg2+, and not further protected by LtrA binding. Importantly, the modification of the A and G residues in the two base pairs above and below A556 in the free RNA implies that the formation of these base pairs and the consequent bulging of A556 must be stabilized by LtrA.
It is also instructive to compare our results with those of the previous in vitro selection, in which DIVa positions 537–598 were "doped" 30%. In that selection, both A556 and G557 were invariant in all tightly binding DIVa variants, whereas other nucleotide residues at the top of stem (ii) as drawn here were conserved in most clones but not invariant (Singh et al. 2002). Twenty-nine of 30 variants could be drawn in a configuration similar to that in Figure 1B with A556 bulged and two base pairs above A556, whereas the remaining variant had just a single GC pair above the bulged A556. There was more variability in the number of base pairs below A556, and in four variants, a G at position 555 was also bulged before resuming stem (ii). Importantly, A556 was unpaired in all of the variants. The greater diversity of structures in the previous selection presumably reflects that more nucleotide residues were allowed to vary, providing more degrees of freedom compatible with LtrA binding.
The binding of LtrA to DIVa is important for translational regulation, RNA splicing, and intron mobility (see Introduction). Our results show that the deletion or pairing of the bulged A556, which is required for tight binding of LtrA to DIV+IVa RNA, had relatively little effect on RNA splicing in vivo (perhaps a slight increase in the level of spliced product), but significantly inhibited intron mobility. Surprisingly, however, the deletion or pairing of the bulged A553, which leads to tighter binding of LtrA, also significantly inhibited mobility. Mutations in the critical nucleotide residues A563 and A564 in the terminal loop (iii), shown previously to inhibit LtrA binding (Singh et al. 2002), inhibited both RNA splicing and intron mobility. Together, these findings show that mutations in different regions of DIVa can differentially affect RNA splicing and intron mobility.
DIVa lies outside the catalytic core, and small mutations in this structure are not expected to strongly affect the ribozyme activity of the intron RNA. The relatively small effects of the DIVa mutations on RNA splicing reflect that LtrA binding to catalytic core regions is more critical for that process than is its binding to DIVa (Cui et al. 2004), whereas the larger effects on intron mobility may reflect that LtrA binding to DIVa is particularly important for positioning the protein for initiation of cDNA synthesis (Wank et al. 1999). Additionally, our results suggest that the binding strength of LtrA to DIVa is delicately tuned, so that either too tight or too weak binding can inhibit mobility. Because LtrA also functions as a translational repressor, too tight binding of LtrA to DIVa could also deleteriously affect translation, particularly if a second round of translation from the same intron is required for synthesis of active LtrA dimers (Cui et al. 2004). These considerations provide an explanation for the persistence of the bulged A553 in the wild-type Ll.LtrB structure, even though its presence impedes LtrA binding.
The LtrA/DIVa interaction characterized here is reminiscent of coat protein/RNA hairpin interactions used for translational repression of viral replicase synthesis in single-stranded RNA phages (Uhlenbeck et al. 1983; Witherell et al. 1991; Draper et al. 1995; Lim and Peabody 2002; Spingola et al. 2002). In these phages, coat protein dimers use a symmetrical ß-sheet formed across the dimer interface to bind an RNA stem–loop that has a bulged A on the 5' side of the stem. The stem–loop contains the replicase initiation region, which is occluded by coat protein binding to down-regulate translation (Draper 1995, 1999). X-ray crystallography of the phage MS2 protein bound to the RNA-stem loop showed that amino acid residues in the ß-sheet recognize specific bases in the loop and the bulged A-residue, which are constrained in a fixed three-dimensional geometry by the stem–loop (Valegard et al. 1994). In other phage, however, the same ß-sheet region has adopted or evolved to recognize divergent stem–loop structures that differ in the size of the loop and stem, and the requirement for bulged nucleotide residues (Draper 1995; Lim and Peabody 2002; Spingola et al. 2002).
The DIVa structures of other group II introns are generally similar to those of the yeast and lactococcal introns in consisting of stem–loops with irregular kinks and bulges (see Singh et al. 2002). The yeast aI1 and aI2 DIVa stem–loops, which are discriminated by their respective IEPs, each contains a 6-nt loop, which differs in sequence between the two introns, and the stems also differ in having bulges at different positions (Huang et al. 2003). Our results suggest that key maturase recognition elements in these and other group II intron DIVas may include the terminal-stem loop, as well as bulged nucleotide residues farther down the stem. However, the findings for the phage coat protein/ RNA hairpin interactions suggest that different IEPs could have diverged to recognize somewhat different elements in this common structural framework.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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was used for cloning. Cells were grown in LB medium, except those for LtrA protein expression, which were grown in SOB medium. Antibiotics were added at the following concentrations: ampicillin (100 µg/mL), chloroamphenicol (25 µg/mL), and tetracycline (25 µg/mL).
Recombinant plasmids
pKW-DIV+IVa and its mutant derivatives, used to synthesize in vitro transcripts for RNA binding experiments, contain the DIV+IVa region of the Ll.LtrB-ORF intron (positions 526–605/ 2383–2395; the discontinuity in position numbers reflects deletion of the intron ORF) cloned behind a phage T7 promoter in pUC19 (Yanisch-Perron et al. 1985). To construct these plasmids, two PCRs were carried out using as template an Ll.LtrB-ORF intron deleted for DIVb (positions 606–2382; Wank et al. 1999). One PCR used the primer DIVupL (5'-AGGAAAGCTTTAATACGACT CACTATAGGGCGCAGTCCAACTCACAGGTTATTGTGTACTA AAATTAAAA), which introduces a HindIII site, T7 promoter, and the 20-nt 5' flanking sequence GGGCGCAGTCCAACTCACAG, in combination with a 30- to 38-nt DNA oligonucleotide primer encoding the mutant DIVa sequence. The other PCR used an overlapping oligonucleotide complementary to the DIVa primer and the primer DIVdnL (5'-TCGTGTGTCTAGACGCGTGAT CAGTTATTGTTCGTTATTCTTTC), which introduces a 10-nt 3' flanking sequence (TGATCACGCG) and an XbaI site. The products of the two PCRs were gel purified, mixed, and amplified with the outside primers pDIVupL and pDIVdnL, and the resulting 162- to 164-bp PCR products, corresponding to the wild-type sequence or mutant sequences with 1-nt deletions or insertions, were gel purified, digested with HindIII and XbaI, and cloned between the corresponding sites of pUC19.
pACD2, the donor plasmid used in intron mobility assays, contains a 0.9-kb Ll.LtrB-ORF intron and short flanking exons cloned behind a T7lac promoter in a CamR pACYC184-derivative (Guo et al. 2000; Karberg et al. 2001). The intron contains an additional T7 promoter inserted near the 3' end of DIV, and the LtrA protein is expressed from a position just downstream from the 3' exon. The recipient plasmid pBRR3-ltrB contains the Ll.LtrB homing site (positions –30 to +15 relative to the intron-insertion site) cloned upstream of a promoterless tetR gene in an AmpR pBR322-derivative (Guo et al. 2000; Karberg et al. 2001). DIVa mutations were recloned into pACD2 from the pKW-DIV+IVa plasmids described above by carrying out PCRs of pACD2 with primers S1 (5'-CACCACATTTGTACAATCTGTAG GAGAACC) plus A1 (5'-CAATTTTTAATTTTAGTACACAATAA CTGTACC) to amplify intron positions 299–552, and with primers S2 (5'-CAATGGCAATTTTAGAAAGAATCAG) plus A2 (5'-CATTGCCATTTCCCAACGCGTCGCCAC) to amplify intron positions 584–690/2287–2301. The gel-purified PCR products were then mixed with the appropriate pKW-DIV+IVa plasmid, and PCR was carried out with primers S1 and A2 to yield a continuous DNA fragment corresponding to Ll.LtrB intron positions 229–690/2287–2301. The final gel-purified PCR product was digested with BsrGI and MluI and cloned between the corresponding sites of pACD2.
pELG2, used for in vivo splicing assays, is an AmpR pET-11a-derivative, which expresses the Ll.LtrB-ORF intron and short flanking exons, with the 3' exon fused in frame to GFP, so that splicing of the intron is linked to the expression of GFP (Cui et al. 2004). The ltrB/GFP fusion is cloned downstream from a T7lac promoter, with the LtrA protein expressed from a position just downstream from the 3' exon/GFP sequence. To reclone DIVa mutations into pELG2, the corresponding pACD2-based plasmid was first digested with SalI and religated to delete the internal T7 promoter in DIV, which would otherwise interfere with the analysis, then digested with HindIII and KpnI to yield a 957-bp fragment that was recloned between the corresponding sites of pELG2.
For each construct, the inserted region was sequenced completely to verify that no adventitious mutations had been introduced during PCR.
Preparation of LtrA protein
The LtrA protein was expressed in E. coli BL21(DE3) from the intein-based expression vector pImp-1P and purified as described (Saldanha et al. 1999). Protein concentrations were determined by Bradford assay (Coomassie Protein Assay Reagent; Pierce), using bovine serum albumin as a standard, with a correction based on a calibrated LtrA preparation whose concentration was determined by A280 (Saldanha et al. 1999). LtrA preparations were checked for in vitro splicing activity before use.
In vitro transcription
In vitro transcription was carried out with phage T7 RNA polymerase (Stratagene) in the manufacturer’s buffer with 2.5 mM of each NTP, 10 mM dithiothreitol, 3.75 mM MgCl2, 0.28 unit/µL RNase inhibitor (Amersham Bioscience) for 3 h at 30°C. To synthesize 32P-labeled RNAs for competition binding assays, the reaction medium was supplemented with 330 nM [-32P]UTP (3,000 Ci/mmole; DuPont-New England Nuclear). Following transcription, the DNA template was digested with DNase I (10 units, 15 min, 37°C; Amersham Bioscience), and transcripts were extracted with phenol-CIA (phenol/chloroform/isoamyl alcohol, 25:24:1 by volume), and ethanol precipitated. Transcripts used for in vitro selection were purified by gel filtration twice through Sephadex G25 (Sigma-Aldrich), and those used in the competition binding assays were purified by electrophoresis in a denaturing 6% polyacrylamide gel.
In vitro selection of DIVa variants
The initial DIVa pool for in vitro selection was generated by PCR of a 136-nt DNA oligonucleotide, randmDIV (5'-GGGCCAGTCC AACTCACAGGTTATTGTGTACTAAAATTAAAAATTGNNNAG NGAGGAAAACCTCNNNNNNNNNNNNNCAATGGCAATTTT AGAAAGAATAACGAACAATAACTGATCACGCGTCTAGACA CACGA; where N = indicates synthesis with equimolar concentrations of G, A, T, and C), with primers DIVup (5'-AGGAAAGC TTTAATACGACTCACTATAGGGCGCAGTCCAACTCACAG) and DIVdn (5'-TCGTGTGTCTAGACGCGTGATCAG). The PCR was carried out in 25 100-µL reactions, each containing 20 pmoles of the DNA oligonucleotide, 100 pmoles of each primer, 0.25 mM of each dNTP, 2.5 units Taq DNA polymerase (Invitrogen), and buffer supplied by the manufacturer. Thermal cycle conditions were seven cycles of 93°C for 30 sec, 55°C for 10 sec, and 72°C for 30 sec, the number of cycles being limited to minimize duplication of DNA sequences. The PCR product was purified in a 1% agarose gel and transcribed with phage T7 RNA polymerase in the presence of [-32P]UTP to generate the initial RNA pool.
For in vitro selection, 200 pmoles of the 32P-labeled RNA pool were dissolved in 20 µL of distilled water and denatured by heating to 90°C for 1 min and then quickly cooled by adding 780 µL prewarmed (30°C) renaturation buffer (128 mM NH4Cl, 6.41 mM MgCl2, 51.3 mM Tris-HCl at pH 7.5, 12.8 mM dithiothreitol, 0.28 unit/µL RNase inhibitor; Amersham Bioscience). After incubating at 30°C for 1 min, 100 µL 4 M NH4Cl was added to bring the final NH4Cl concentration to 500 mM, followed by 100 µL LtrA protein (1.5 µM for the initial round and 0.1 µM for subsequent rounds). The solution was incubated for 1 h at 30°C, then filtered through a nitrocellulose membrane filter (BA85; Schleicher & Schuell) to collect RNA–protein complexes. After washing three times with 2 mL AMT (500 mM NH4Cl, 5 mM MgCl2, 40 mM Tris-HCl at pH 7.5), RNA was extracted by vortexing the filter in 400 µL 10 mM Tris-HCl (pH 7.5), 1 mM EDTA plus 400 µL phenol-CIA and then incubating for 3 h at room temperature. The extracted RNA was ethanol precipitated and reverse transcribed using Superscript II RT (Invitrogen) with primer DIVdn, following the manufacturer’s protocol, then PCR amplified with primers DIVup and DIVdn to generate the DNA pool for the next round of selection.
Competition binding assays
Competition binding assays were carried out as described (Saldanha et al. 1996). 32P-labeled wild-type and mutant RNAs were transcribed from cloned (individual variants) or PCR-generated DNA templates (RNA pools) and tested against a wild-type DIV+IVa competitor RNA transcribed from a clone with shorter 5'- and 3'-flanking sequences (GGG and TCTAG, respectively). A mixture containing 100 nM of each RNA was incubated with 10 nM LtrA protein in 100 µL AMT containing 5 mM dithiothreitol and 0.28 unit/µL RNase inhibitor (Amersham Bioscience) for 60 min at 30°C, and then filtered through nitrocellulose to bind RNA–protein complexes. The RNAs were recovered from the filter by phenol-CIA extraction, as described above, followed by ethanol precipitation in the presence of 1/10 volume of 3 M ammonium acetate and E. coli tRNA carrier (10–20 µg; Sigma-Aldrich), and then analyzed in a denaturing 5% polyacrylamide gel, which was dried and quantified using a PhosphorImager (model 445SI; Molecular Dynamics). The binding ratio is defined as the ratio of the mutant to wild-type RNA bound by LtrA relative to the input ratio of the RNAs. To control for the effect of different flanking sequences used to generate different-sized transcripts, the binding ratios were normalized based on binding ratios determined in parallel for two wild-type RNAs having the same flanking sequences (1.17 and 1.19 in two experiments).
FACS assay of in vivo RNA splicing
E. coli HMS174(DE3), which carries an IPTG-inducible T7 RNA polymerase, was transformed with pELG2 plasmids containing wild-type and mutant Ll.LtrB introns, and a single colony was inoculated into 5 mL LB medium plus 100 µg/mL ampicillin and grown overnight with shaking at 37°C. Fifty microliters of the overnight culture were then inoculated into 5 mL of flesh LB plus ampicillin, grown at 37°C until O.D.595 = 0.4, and induced with 100 µM IPTG for 3 h at 37°C. Cells were assayed for GFP fluorescence by using a fluorescence-activated cell sorter (FACS Caliber; Becton Dickinson, Immunocytometry Systems), with filter FL1 (530 ± 30 nm). The data were analyzed by using the CELLQuest program (Becton Dickinson).
Poisoned primer extension assay of in vivo splicing
Poisoned primer extension assays (Pikielny and Rosbash 1985) were carried out with whole-cell RNA from E. coli HMS174(DE3) containing pELG2-WT or DIVa mutants and grown and induced with IPTG, as described above for FACS assay. The RNA was extracted as described by Belfort et al. (1990) and further purified by gel filtration (Centri-Sep; Princeton Separations). Five micrograms RNA were annealed with 0.5 pmoles 5' end-labeled primer 5'-GATTCGTAGAATTAAAAATGATATGGT by incubating in 7 µL 50 mM Tris-HCl (pH 7.5), 60 mM NaCl, 10 mM dithiothreitol for 3 min at 60°C, then chilling on ice. After additions to give 125 µM each dATP, dTTP, and dGTP, 500 µM ddCTP, and 1.8 mM MgCl2, the reaction was initiated by adding 180 units M-MLV RT (Invitrogen), incubated for 30 min at 48°C, and terminated by adding gel loading buffer (80% formamide, 10 mM EDTA at pH 8.0, xylene cyanol [1 mg/mL], and bromophenol blue [1 mg/mL]). The products were analyzed in a denaturing 10% polyacrylamide gel, which was dried and quantified with a PhosphorImager.
Intron mobility assays
Intron mobility frequencies were determined by using an E. coli two-plasmid assay in which a 0.9-kb Ll.LtrB-ORF intron with a phage T7 promoter inserted near its 3' end is expressed from a donor plasmid and integrates into a recipient plasmid target site cloned upstream of a promoterless tetR gene, thereby activating that gene (Guo et al. 2000; Karberg et al. 2001). The CapR intron donor plasmid pACD2 and the AmpR recipient plasmid pBRR3-ltrB were cotransformed into E. coli HMS174(DE3), and the cells were grown and induced with 100 µM IPTG for 1 h at 37°C, as described (Guo et al. 2000; Karberg et al. 2001), then plated on LB medium containing ampicillin or ampicillin plus tetracycline. Mobility frequencies are defined as the ratio of (TetR + AmpR)/AmpR colonies.
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ACKNOWLEDGMENTS |
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The publication costs of this article were defrayed in part by payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 USC section 1734 solely to indicate this fact.
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Footnotes |
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Received April 29, 2004; accepted June 7, 2004.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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