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Nucleic Acids Research, 2003, Vol. 31, No. 9 2381-2392
© 2003 Oxford University Press

RNA structure-dependent uncoupling of substrate recognition and cleavage by Escherichia coli ribonuclease III

Irina Calin-Jageman and Allen W. Nicholson

Department of Biological Sciences, Wayne State University, Detroit, MI 48202, USA

*To whom correspondence should be addressed at present address: Department of Chemistry, Temple University, 1901 N. 13th Street, Philadelphia, PA 19122, USA. Tel: +1 215 204 4410; Fax: +1 215 204 1532; Email: anichol{at}temple.edu

Received January 6, 2003; Revised and Accepted March 6, 2003


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Members of the ribonuclease III superfamily of double-strand-specific endoribonucleases participate in diverse RNA maturation and decay pathways. Ribonuclease III of the gram-negative bacterium Escherichia coli processes rRNA and mRNA precursors, and its catalytic action can regulate gene expression by controlling mRNA translation and stability. It has been proposed that E.coli RNase III can function in a non-catalytic manner, by binding RNA without cleaving phosphodiesters. However, there has been no direct evidence for this mode of action. We describe here an RNA, derived from the T7 phage R1.1 RNase III substrate, that is resistant to cleavage in vitro by E.coli RNase III but retains comparable binding affinity. R1.1[CL3B] RNA is recognized by RNase III in the same manner as R1.1 RNA, as revealed by the similar inhibitory effects of a specific mutation in both substrates. Structure-probing assays and Mfold analysis indicate that R1.1[CL3B] RNA possesses a bulge– helix–bulge motif in place of the R1.1 asymmetric internal loop. The presence of both bulges is required for uncoupling. The bulge–helix–bulge motif acts as a ‘catalytic’ antideterminant, which is distinct from recognition antideterminants, which inhibit RNase III binding.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Members of the ribonuclease III superfamily of double-stranded (ds) RNA-specific endoribonucleases play essential roles in prokaryotic and eukaryotic RNA maturation and decay pathways (1). Eukaryotic RNase III orthologs participate in ribosomal RNA maturation and cleave precursors to snRNAs and snoRNAs (1,2). The functionally and structurally distinct eukaryotic ortholog Dicer performs a critical early step in RNA interference (RNAi), by cleaving dsRNAs to 21–23 bp fragments. These species, termed small interfering (si) RNAs, exert selective inhibition of gene expression through homology-dependent RNA degradation (3–5). Dicer also cleaves precursors to micro-RNAs, which exert cistron-specific translational control, and perhaps participate as well in other gene regulatory mechanisms (6,7).

RNase III orthologs are highly conserved in the Bacteria, and participate in species-specific RNA maturation and decay pathways as well as in rRNA processing (8,9). Bacterial RNase III orthologs exhibit the simplest primary structures of the superfamily members, and consist of a C-terminal dsRNA-binding motif (dsRBM) (10,11) and an N-terminal catalytic (nuclease) domain (1,12–15). The most-studied Bacterial ortholog is Escherichia coli RNase III (1,8,9,16), which is active as a homodimer, and requires a divalent metal ion (preferably Mg2+) to hydrolyze phosphodiesters. The dsRBM and catalytic domains as isolated polypeptides possess dsRNA-binding and dsRNA-cleaving activities, respectively (17). The catalytic domain exhibits the same strict dsRNA specificity and dimeric structure as the holoenzyme (17). Thus, the dsRBM is not required for conferring double-strand specificity nor is it critical for dimer stability. These findings are consistent with the crystal structure of the catalytic domain of Aquifex aeolicus RNase III, which is dimeric and exhibits an extensive subunit interface that defines a putative dsRNA-binding cleft (15).

Escherichia coli RNase III recognizes its substrates through specific structural and sequence features (reactivity epitopes) that are contained within a double helical structure of at least one full turn (>11 bp) (8,9,18). Two specific W-C base-paired regions, termed the proximal box (pb) and distal box (db) represent sites of enzyme–substrate contacts (19) as well as sites in which specific W-C bp inhibit RNase III binding (20). The inhibitory W-C bp are termed RNase III antideterminants, and are proposed to play a role in cleavage site selection as well as protect other dsRNAs with important functions from inadvertent cleavage (9,20).

RNA internal loops represent an additional type of reactivity epitope that can alter the normal pattern of double-strand processing, to cleavage of a single strand. Bacteriophage T7 expresses transcripts containing RNase III cleavage sites. The T7 substrates are hairpin structures with internal loops, and cleavage of a single phosphodiester within the internal loop 3' segment separates the flanking mRNAs, allowing their independent translation. The prolonged physical half-lives of T7 mRNAs are also due in part to the 3' hairpin structures created by the catalytic action of RNase III (21). One of the T7 substrates is R1.1 RNA (Fig. 1A, inset), which has been the subject of a number of biochemical and structural studies to identify substrate reactivity epitopes (20,22–25).




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  		Figure 1. (Previous page and above) In vitro selection strategy for isolating cleavage-resistant variants of R1.1 RNA. (A) Structure of R1.1[SxN] RNA. The nine sequence-randomized sites (N) in the internal loop are indicated. The inset figure shows the sequence of R1.1 RNA, with the primary (1°) and secondary cleavage sites indicated by the solid and dashed arrows, respectively. The positions of the SacI and XhoI restriction sites are indicated, as are the positions and sequences of the forward PCR primer and reverse transcriptase (RT) primer. (B) In vitro selection strategy. The oligodeoxynucleotide encoding R1.1[SxN] RNA is transcribed to provide an RNA pool, which then is incubated with RNase III. The products are reversed transcribed and the cDNAs subjected to PCR with the primers shown in (A). Only cDNAs corresponding to uncleaved RNA sequences can be amplified, providing a template pool for a new round of transcription and selection. The final amplified DNA sequences are cleaved by XhoI and SacI, and cloned into a pBluescript plasmid. See Materials and Methods for additional information.

 
Escherichia coli RNase III also cleaves transcripts expressed by bacteriophage lambda, and is an important participant in the lysis/lysogeny decision (8,26). Translation of the lambda cIII mRNA is RNase III-dependent (27), and it has been proposed that cIII protein synthesis is stimulated by binding of RNase III to the cIII mRNA 5' leader sequence without concomitant cleavage (27). However, there has been no direct biochemical evidence for such a function of E.coli RNase III, or any other RNase III ortholog. Given the key functional roles of RNase III superfamily members in global regulation, host defense and genome maintenance, it is of interest to determine whether RNA structures can be identified that allow binding of RNase III, but are resistant to cleavage. We describe in this report the use of in vitro selection technology to isolate and characterize a binding-competent, cleavage-resistant RNA ligand for RNase III.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Materials
Chemicals and reagents were molecular biology grade and were generally obtained from Sigma or Fisher Scientific. Water was deionized and distilled. Ribonucleoside 5'-triphosphates were purchased from Amersham-Pharmacia Biotech. [{gamma}-32P]ATP (3000 Ci/mmol) and [{alpha}-32P]CTP or [{alpha}-32P]UTP (3000 Ci/mmol) were from Perkin-Elmer. Escherichia coli bulk stripped tRNA was purchased from Sigma and further purified by repeated phenol extraction and ethanol precipitation. Restriction enzymes and T4 polynucleotide kinase were purchased from New England Biolabs and used with the supplied buffers. Calf intestine alkaline phosphatase was obtained from Roche Molecular Biochemicals. T7 RNA polymerase was purified from an overproducing E.coli strain as described (28). (His)6-RNase III was purified as described (29). Oligodeoxynucleotide transcription templates were synthesized by Invitrogen or by Midland Certified Reagent Company and were further purified by denaturing gel electrophoresis (29). Purified DNAs were stored at –80°C in Tris–EDTA buffer (pH 8).

RNA synthesis
Oligodeoxynucleotide-directed transcription reactions were performed using T7 RNA polymerase essentially as described (29,30). For synthesis of internally 32P-labeled RNA, transcription reactions (100 µl) included the four rNTPs (1 mM each), ~3–15 µCi [{alpha}-32P]UTP or [{alpha}-32P]CTP, and ~400 U of T7 RNA polymerase. Reactions were incubated at 37°C for 3–4 h. For 5' 32P-labeling, unlabeled purified transcripts were dephosphorylated using calf intestine alkaline phosphatase, then treated with 3–10 µCi [{gamma}-32P]ATP (3000 Ci/mmol) and T4 polynucleotide kinase (2 U). RNAs were purified by denaturing polyacrylamide gel electrophoresis (29) and stored in TE buffer at –20°C.

Substrate cleavage assay
RNase III cleavage assays were performed essentially as described (29). To remove intermolecular complexes formed upon storage at –20°C, 32P-labeled RNA in TE buffer was heated at 100°C for 30 s, then placed on ice. An aliquot of the RNA was added to a reaction containing: 250 mM potassium glutamate (or 160 mM NaCl), 30 mM Tris–HCl (pH 8), 0.01 µg/µl tRNA, 0.1 mM EDTA and 0.1 mM DTT. In some reactions, 5% glycerol (v/v) was included. RNase III was added at the specified concentration and cleavage reactions initiated by adding MgCl2 (10 mM final concentration). Reactions were incubated at 37°C and stopped by addition of an equal volume of gel loading buffer containing 20 mM EDTA and 7 M urea (29). Aliquots were electrophoresed (25 V/cm) for 1–2 h in a 15% polyacrylamide gel containing 7 M urea and TBE buffer. Reaction products were visualized by phosphorimaging (Molecular Dynamics Storm 860 system), and quantitation performed (29) using ImageQuant software.

Gel mobility shift assay
Gel shift assays were performed essentially as described (29). To remove intermolecular complexes formed on storage at –20°C, 5' 32P-labeled RNA was heated in TE buffer in a boiling water bath for 30 s, then immediately cooled on ice. An aliquot of the RNA (~8000 d.p.m., ~1–2 fmol) was combined with RNase III in binding buffer [160 mM NaCl, 10 mM MgCl2 (or 10 mM CaCl2), 30 mM Tris–HCl (pH 8), 0.01 µg/µl tRNA, 0.1 mM EDTA, 0.1 mM DTT, 5% glycerol]. Samples were incubated at 37°C for 10 min, placed on ice for 20 minutes, then electrophoresed (120 V at 4°C for 3 h) in a 7% polyacrylamide gel (80:1 acrylamide:bisacrylamide) containing TBE buffer supplemented with 10 mM MgCl2 (or 10 mM CaCl2). Binding reactions were visualized and quantitated as described (29).

In vitro selection procedure
Oligodeoxynucleotides (128 nt) encoding R1.1[Sx] RNA or R1.1[SxN] RNA (see Fig. 1A) (sequences available upon request) were synthesized by Midland Certified Reagent Company and were purified by gel electrophoresis (29). RNA was synthesized by T7 RNA polymerase transcription of the oligonucleotides using [{alpha}-32P]CTP as the radiolabel (see above). Gel-purified RNA (~15 000 d.p.m., 5.4 pmol) was incubated with RNase III (200 nM concentration) for 40 min at 37°C. An aliquot was analyzed by gel electrophoresis to assess extent of cleavage (see above). The remainder of the reaction (see above) was treated with 4 U of RNase-free DNase I (Ambion) at 37°C for 40 min, which was then inactivated by heating at 75°C for 10 min. An aliquot of the DNase-treated RNA (~1.1 pmol RNA) was combined with reverse transcriptase (RT) primer (2 pmol) (see Fig. 2A) in 10 µl of water. The sample was heated at 70°C for 10 min, cooled to room temperature, then placed on ice. The sample was supplemented with first-strand cDNA synthesis buffer (Superscript II system; Invitrogen) which included 10 mM DTT and 0.5 mM each dNTP. The sample was incubated at 42°C for 2 min, Superscript II Reverse Transcriptase (Invitrogen, 400 U) added, and the reaction (20 µl final volume) incubated at 42°C for 50 min, followed by 70°C for 15 min. Reactions were stored at –20°C prior to further manipulation.




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  		Figure 2. (Previous page and above) RNase III cleavage reactivity of R1.1[SxN] RNA as a function of selection round. (A) Cleavage patterns of R1.1[Sx] RNA, R1.1[SxN] RNA (initial pool) and R1.1[SxN] RNA (round 6). Lanes 1, 3 and 5 show reactions incubated for 40 min in the presence of RNase III (200 nM), but without MgCl2. Lanes 2, 4 and 6 show reactions incubated for 40 min with RNase III in the presence of MgCl2. Products were electrophoresed in a 15% polyacrylamide gel containing 7 M urea and were visualized by phosphorimaging. To highlight the cleavage-resistance the amounts of RNA analyzed in lanes 5 and 6 were greater than in the other lanes. The arrow marked ‘1°’ indicates the position of two fragments of approximately equal lengths (53 and 56 nt), created by cleavage of the 109 nt RNA at the primary site (shown in Fig. 2A). ‘US’ and 5' indicate the additional products of cleavage (both 28 nt in size) at the secondary site. The species indicated by arrowheads indicate products of cleavage at unidentified sites. (B) Resistance of R1.1[SxN] RNA to RNase III cleavage as a function of selection round. Aliquots of 32P-labeled RNA transcribed from the amplified DNA after each round were incubated with RNase III, then electrophoresed as described above. Percent resistance to cleavage was determined by phosphorimaging (see Materials and Methods).

 
For PCR amplification, one-tenth of the RT reaction was combined with 30 pmol of RT primer and 30 pmol of T7 primer (see Fig. 1A) in 50 µl of the supplied PCR buffer, which also contained 1.5 mM MgCl2, 0.2 mM dNTP mix and 2.5 U of Amplitaq DNA polymerase (Perkin-Elmer). The thermal cycler settings were: 94°C for 3 min (initial cycle only); 94°C for 15 s; 55°C for 30 s; and 72°C for 30 s (32 cycles total). The PCR products were gel purified using a Qiaquick MinElute kit (Qiagen) or a Zymo-Clean kit (Zymo Research, Orange, CA). T7 RNA polymerase transcription (see above) provided substrate for a new round of selection.

For plasmid cloning of PCR products, ~1.5 µg of the PCR-amplified DNA was treated with SacI (20 U) at 37°C for 1 h. An additional 20 U of SacI was added and the reaction incubated for 1 h at 37°C, then at 65°C for 20 min. XhoI (20 U) was added and the reaction incubated at 37°C followed by addition of a second, equal amount of enzyme and incubation for 1 h at 37°C. The reaction was stopped by heating at 65°C for 15 min. The DNA was purified using a Qiaquick MinElute kit and stored at –20°C in TE buffer prior to further use. An aliquot of purified, digested DNA (see above) was ligated to XhoI, SacI-digested plasmid pBSII(KS–) (Stratagene) using T4 DNA ligase. Transformation of E.coli DH10B cells was carried out by electroporation. Ampicillin-resistant bacterial colonies were isolated, and plasmids purified and sequenced.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
In vitro selection of cleavage-resistant variants of T7 R1.1 RNA
R1.1 RNA (Fig. 1A, inset) is a ~60 nt hairpin containing an asymmetric [4nt/5nt] internal loop, and is located in the T7 genetic early region between genes 1 and 1.1. R1.1 RNA is cleaved by RNase III at a single phosphodiester (indicated by the arrow in Fig. 1A, inset), which directly creates the mature 3' and 5' ends of the flanking gene 1 and 1.1 mRNAs, respectively (21). Two recent findings prompted the use of R1.1 RNA as a starting substrate to search for a cleavage-resistant, binding-competent sequence. First, an R1.1 RNA variant which contains a sequence-randomized internal loop retains comparable cleavage reactivity as the parent substrate (I.C.-J. and A.W.N., manuscript submitted for publication). While this finding demonstrated the absence of a strict sequence requirement for cleavage, it was also noted that ~10% of the sequence-randomized R1.1 RNA pool was highly resistant to cleavage. This raised the possibility that some of the cleavage-resistant sequences might possess appreciable binding affinity. Secondly, the intercalating agent ethidium bromide can uncouple E.coli RNase III binding and cleavage of R1.1 RNA in vitro (31). The uncoupling is dependent upon formation of an ethidium–substrate complex which retains binding affinity but is refractory to cleavage (31). An ethidium binding site implicated in inhibition was mapped to the internal loop (31). Thus, a ligand-induced alteration in internal loop structure can suppress the catalytic step, raising the possibility that a specific RNA structure may exist which could exert the same effect.

We used in vitro selection (SELEX) (32–35), and specifically applied a negative selection strategy wherein only RNA sequences resistant to cleavage by RNase III would be competent for amplification by reverse transcription and PCR (RT–PCR) (Fig. 1B) using the primer pair shown in Figure 1A. The cleavage-resistant population would be expected to include any binding-competent, cleavage-resistant sequences, which would be identified in a subsequent screen. The starting substrate was R1.1[SxN] RNA (Fig. 1A), a 109 nt R1.1 RNA variant containing a randomized internal loop of 49 (262 144) different sequences. To assist in amplification, cloning and sequence analysis, the 5' and 3' flanking sequences of R1.1[SxN] RNA contained RT and PCR primer binding sites, as well as restriction enzyme sites (Fig. 1A).

The cleavage reactivity of R1.1[SxN] RNA was compared with R1.1[Sx] RNA, which contains the R1.1 internal loop sequence. Cleavage of internally 32P-labeled R1.1[Sx] RNA at the canonical site is expected to produce two radiolabeled fragments of approximately equal size (53 and 56 nt). The formation of these products is consistent with a denaturing polyacrylamide gel analysis of a representative in vitro cleavage reaction (Fig. 2A). The assay shows that R1.1[Sx] RNA and R1.1[SxN] RNA are both reactive, with the single predominant band (Fig. 2A, lanes 2 and 4) consistent with the RNase III-dependent production of two similarly sized species. Although this gel system was unable to resolve the two products, the relevant result is that the cleavage pattern of R1.1[SxN] RNA is qualitatively the same as that of R1.1[Sx] RNA. The shorter fragments produced in lower amounts (indicated by the arrowheads in Fig. 2A) include the products of cleavage of the secondary site in the internal loop (indicated in Fig. 1A). Secondary sites are recognized at higher enzyme concentrations (36). In summary, the similar cleavage patterns of the two RNAs is consistent with the conservation of the canonical cleavage site in many other R1.1 RNA variants (22–24), and justifies the use of R1.1[SxN] RNA as a substrate for in vitro selection.

To maximize cleavage of the reactive population, R1.1[SxN] RNA was treated with a high (200 nM) concentration of RNase III. The products were subjected to RT–PCR and the amplified DNA sequences transcribed to provide substrate for a new round of selection and amplification. Aliquots of the 32P-labeled transcripts from each round were assessed for their resistance to cleavage. Figure 2B summarizes the cleavage assays, and shows that the early rounds provided an R1.1[SxN] RNA population exhibiting a successively greater resistance to cleavage. By round four, the percent resistance to cleavage approached 80%, which was essentially unchanged by round six (Fig. 3B). The ~20% residual reactivity probably reflects several factors, including forcing conditions for enzymatic cleavage, the occurrence of nonspecific (i.e. nonenzymatic) cleavage events and new sequence heterogeneity generated during each round of transcription and RT–PCR.



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  		Figure 3. Cleavage resistance of Class I R1.1 RNA variants, and involvement of the A9U mutation. (A) Analysis of R1.1[CL68B] RNA. The sequence of the RNA is shown on the left. The right side presents a cleavage assay showing the resistance of internally 32P-labeled R1.1[CL68B] RNA to RNase III (lanes 1 and 2), and restoration of reactivity by the U9A reversion (lanes 3 and 4). The reactions involved incubation with 10 nM enzyme for 40 min. The four bands in lane 4 include the products of cleavage at the primary site (indicated by 1° and 3'), or at the primary and secondary sites (indicated by US and 5'; US, upper stem). (B) Analysis of R1.1[CL14A] RNA reactivity. The left side shows the structure of R1.1[CL14A] RNA. The assay on the right shows the cleavage resistance of internally 32P-labeled substrate (compare lanes 1 and 2) and restoration of reactivity by the U9A reversion (lanes 3 and 4). The reaction conditions were the same as the experiment in (A).

 
To assess general trends in the sequence of the internal loop, batch sequencing of the PCR products was performed after each round. The results (data not shown) revealed an enrichment in cytosine residues at most of the positions. This trend was verified by sequencing individual clones (see below). The R1.1[SxN] RNA pool obtained at round six was subjected to RT–PCR, the amplified DNAs cloned into a plasmid, and the sequences determined for 330 clones. Table 1 provides the aligned sequences of 86 of the 330 cloned sequences. The 86 sequences were designated either as Class I or Class II R1.1 RNA variants (see below), and were chosen on the basis of the absence of nucleotide deletions, insertions or other undirected mutations external to the internal loop (with one important exception, as described below). Approximately 40 of the sequenced plasmids were linearized by XhoI digestion, the cloned sequence transcribed in vitro by T7 RNA polymerase, and the RNAs assessed for their RNase III cleavage reactivities. The results (data not shown) reveal that the selected R1.1 sequences are resistant to cleavage.


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  		Table 1. Internal loop sequences of R1.1[SxN] RNAs selected after round six
 
Analysis of Class I R1.1 RNA variants
The Class I R1.1[Sx] RNA variants exhibit cytosine-rich internal loops. The internal loop 5' segment (positions 17–20) exhibits a >50% average cytosine content at each of the four positions (Table 1). Moreover, 72 of the 78 Class I RNAs contain two or three adjacent cytosine residues within the internal loop 3' segment. The cytosine enrichment is consistent with the batch DNA sequencing analysis. The Class I R1.1 RNA variants also exhibit a single nucleotide change (A9U) in the lower stem, which changes the canonical AU W-C base pair to a UU mismatch. Batch DNA sequence analysis (data not shown) revealed that this mutation dominates the R1.1[SxN] RNA pool after the fourth round of selection. The involvement of this unprogrammed mutation in conferring cleavage resistance is addressed below.

The reactivities of the Class I R1.1[Sx] RNA variants were examined within the context of the 60 nt R1.1 RNA structure. The internal loop sequences and the A9U mutation of clones R1.1[CL68B] and R1.1[CL14A] were incorporated into R1.1 RNA, providing R1.1[CL68B] RNA and R1.1[CL14A] RNA. To assess the involvement of the A9U mutation, the same R1.1 RNA variants were prepared but which instead lacked this mutation (R1.1[CL68B;U9A] RNA and R1.1[CL14A;U9A] RNA). A cleavage assay shows that R1.1[CL68B] RNA and R1.1[CL14A] RNA are resistant to RNase III (Fig. 3A, lane 2 and B, lane 2). Thus, the inhibitory actions of the internal loop sequences are manifested in the 60 nt canonical R1.1 hairpin. Gel shift assays (data not shown) revealed that neither R1.1[CL68B] RNA nor R1.1[CL14A] RNA are detectably bound by RNase III. The same variants which instead lack the A9U mutation are cleaved by RNase III (Fig. 3A, lane 4 and Fig. 3B, lane 4). However, the A9U mutation on its own does not inhibit cleavage of either R1.1 RNA or R1.1[Sx] RNA (data not shown). We conclude that the A9U mutation cooperates with the variant R1.1 internal loop sequences to confer inhibition.

The cleavage resistance of the Class I R1.1 RNA variants may reflect formation of a novel RNA conformation. To examine this we used native gel electrophoresis, which can detect alternative RNA conformations (37). The Class I R1.1 RNA variants R1.1[CL68B] RNA and R1.1[CL14A] RNA, as well as their counterparts lacking the A9U mutation were electrophoresed in a nondenaturing polyacrylamide gel. The three cleavage-resistant Class I RNAs electrophorese at a faster rate than R1.1 RNA, while the cleavage-competent RNAs lacking the A9U mutation comigrate with R1.1 RNA (data not shown). We conclude that the lack of reactivity of Class I RNAs correlates with an altered conformation, which is in turn dependent upon the A9U mutation. A possible structure of the alternative conformation is considered below (see Discussion).

Analysis of Class II R1.1 RNA variants
The Class II R1.1 RNAs are fewer in number than the Class I RNAs, and lack the A9U mutation (Table 1). Cleavage assays were performed on eight Class II R1.1 RNA variants. The results reveal that all of the RNAs are resistant to RNase III (data not shown). Thus, the A9U mutation is not absolutely required for inhibition. A representative variant, R1.1[CL17C] RNA, electrophoreses in a nondenaturing gel at a slightly faster rate than R1.1 RNA (data not shown). Thus, for at least one Class II RNA, cleavage resistance correlates with an altered conformation.

R1.1[CL3B] RNA: a binding-competent, cleavage-resistant Class II R1.1 RNA variant
Gel shift assays revealed that none of the Class II R1.1 RNA variants are detectably bound by RNase III (data not shown), with one exception. A cleavage assay of R1.1[CL3B] RNA (Fig. 4A) confirmed the expected resistance to cleavage by RNase III (Fig. 4B, lanes 7–12). Less than 5% of the RNA is cleaved in the presence of an excess (200 nM) of RNase III. However, a gel shift assay reveals that R1.1[CL3B] RNA binds RNase III to form a complex with similar electrophoretic mobility as the complex involving R1.1 RNA (Fig. 4C, compare lanes 7–12 with lanes 1–6). The complex is detectable at a relatively low (~5 nM) RNase III concentration, with the RNA fully bound by ~50 nM RNase III. The KD for the complex containing R1.1[CL3B] RNA is 43 nM, while the KD of the complex containing R1.1 RNA is 34 nM. While the gel shift assays yielded some variation in KD, the ratio of KD values for the two complexes was consistently 1.5–2.0. We conclude that R1.1[CL3B] RNA binds RNase III with comparable affinity as R1.1 RNA. The gel shift assay also shows that unbound R1.1[CL3B] RNA exhibits a comparable gel electrophoretic mobility as R1.1 RNA (Fig. 4C, compare lanes 1 and 7), indicating similar structures.



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  		Figure 4. Analysis of the Class II RNA, R1.1[CL3B] RNA. (A) The sequence of R1.1[CL3B] RNA. The nucleotide changes in the internal loop are indicated by bold font. The sequence of the R1.1 RNA internal loop is shown in the inset. Also shown is the distal box (db) and the UG->GU ‘db17’ mutation used in the experiment shown in Figure 5. (B) Cleavage assay of R1.1[CL3B] RNA (lanes 7–12) and R1.1 RNA (lanes 1–6) as a function of RNase III concentration. Lanes 1 and 7 represent incubation of RNA for 10 min in the absence of RNase III. Lanes 2–6 and lanes 8–12 show 10 min reactions using RNase III at a concentration of 2, 5, 8, 50 and 100 nM, respectively. The small amount of cleavage of R1.1[CL3B] RNA observed in lane 12 corresponds to 4.6% conversion to product, with cleavage occurring at the canonical cleavage site, based on similar gel electrophoretic mobilities in a denaturing gel (20). (C) Gel shift assay of R1.1[CL3B] RNA (lanes 1–6) and R1.1 RNA (lanes 7–12). Shift assays were performed using RNase III in the presence of Ca2+, which enhances substrate binding but does not support catalysis (see Materials and Methods). Lanes 1 and 7 show the mobility of the two RNAs in the absence of added RNase III. Lanes 2–6 and 7–12 represent RNase III concentrations of 2, 5, 8, 50 and 100 nM, respectively.

 
The comparable binding affinities of R1.1[CL3B] RNA and R1.1 RNA suggest a common mode of recognition by RNase III. However, it is possible that R1.1[CL3B] RNA binds RNase III in a manner distinct from that of a competent substrate. For example, the RNA may bind to a site that does not involve the catalytic site. Since technical limitations prevented the use of competition gel shift or cleavage assays as definitive tests (I.C.-J. and A.W.N., unpublished experiments), we instead examined the effect on RNase III binding of site-specific substrate mutation. The distal box (see Fig. 4A) is a 2 bp segment which engages in contacts with RNase III (19), and is also a site where specific W-C base pair substitutions block RNase III recognition (20). If R1.1[CL3B] RNA and R1.1 RNA are recognized in the same manner, then a distal box mutation that inhibits R1.1 RNA binding should also affect binding of R1.1[CL3B] RNA. A single substitution (UG->GU) was introduced in the lower position of the distal box of R1.1[CL3B] RNA (see Fig. 4A). This substitution has been shown to inhibit RNase III recognition of a minimal variant of R1.1 RNA (A. Pertzev and A.W.N., manuscript in preparation). A gel shift assay reveals that the mutation strongly inhibits R1.1[CL3B] RNA binding to RNase III (Fig. 5A, compare lanes 4–6 with lanes 10–12). As expected, the binding of R1.1 RNA is also inhibited, although to a lesser extent (Fig. 5A, compare lanes 1–3 with lanes 7–9). Quantitation of the gel shift assay is summarized in Figure 5B. The weaker effect of the mutation on R1.1 RNA binding may reflect the somewhat greater binding affinity of this RNA. The common inhibitory effect of a distal box mutation provides strong evidence that R1.1[CL3B] RNA binds RNase III in the same manner as R1.1 RNA.




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  		Figure 5. Common mechanism of RNase III recognition of R1.1[CL3B] RNA and R1.1 RNA. (A) Gel shift assays were performed using 5' 32P- labeled RNA, and RNase III in Ca2+-containing buffer (see Materials and Methods). Lanes 1–3, R1.1 RNA. Lanes 4–6, R1.1[CL3B] RNA. Lanes 7–9, R1.1[db17] RNA. Lanes 10–12, R1.1[CL3B;db17] RNA. See Figure 4A for the location of the db17 mutation. Lanes 1, 4, 7 and 10 show RNA mobility in the absence of RNase III. The paired lanes 2 and 3, 5 and 6, 8 and 9, and 11 and 12 show the mobilities of each RNA in the presence of 50 or 100 nM RNase III, respectively. The positions of the free and bound RNAs are indicated. (B) Graphic presentation of the effect of the db17 mutation on RNase III binding to R1.1[CL3B] RNA and R1.1 RNA. The graph shows the percent RNA bound, as determined by the ratio of the amount of bound RNA to the total (bound plus unbound) RNA. Two RNase III concentrations used were 50 and 100 nM. Shown to the right is the key to the RNAs examined. The binding of R1.1[CL3B; db17] RNA was undetectable.

 
Structural analysis of R1.1[CL3B] RNA
Mfold analysis reveals a hairpin structure for R1.1[CL3B] RNA essentially the same as that of R1.1 RNA. However, the Mfold structure of R1.1[CL3B] RNA has a bulge–helix–bulge motif in place of the R1.1 internal loop (Fig. 6B). Nucleotide A20, and nucleotides A48 and U49 provide 1 and 2 nt bulges, respectively, with the two bulges separated by three GC base pairs (Fig. 6B). Terbium (Tb3+) ion was used as a structural probe of the proposed bulge–helix–bulge motif, and to identify other features potentially involved in uncoupling. Tb3+ cleaves RNA within unstructured, conformationally flexible regions, and is relatively insensitive to specific sequence (38,39). Figure 6A shows the Tb3+ cleavage pattern for 5' 32P-labeled R1.1[CL3B] RNA. For comparative purposes, R1.1 RNA and R1.1[WC-R] RNA were also examined. The latter RNA is a variant of R1.1 RNA in which the internal loop is replaced by a fully W-C base paired sequence (24). For R1.1[CL3B] RNA, Tb3+ cleavage sites occur at A20 and at A48/U49, with some cleavage occurring at the nucleotides that flank the two bulge positions (Fig. 6A, lane 4; summarized in Fig. 6B). The Tb3+ reactivity pattern is consistent with the proposed bulge–helix–bulge motif.




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  		Figure 6. Structural analysis of R1.1[CL3B] RNA. (A) Tb3+ ion structure probing. Terbium ion (Tb3+)-dependent RNA cleavage was carried out essentially as described (38,39) with some modification. TbCl3 was dissolved in 5 mM HEPES (pH 5.5) at a final concentration of 0.5 M and stored at –20°C prior to use. Briefly, 5' 32P-labeled RNA (~15 000–20 000 c.p.m.) was incubated with 5 mM TbCl3 in 20 mM NaCl, 50 mM HEPES (pH 7.5) in a 10 µl reaction for 15 min at 37°C. The reaction was stopped by EDTA (2 µl of a 250 mM solution). Samples were ethanol precipitated and resuspended in a small volume of TE buffer. An equal volume of deionized formamide (containing 0.025% bromophenol blue) was added, and the sample electrophoresed (1500 V, 2–4 h) in a 10% polyacrylamide gel containing 7M urea. The reactions were visualized by phosphorimaging. Lanes 1–3, R1.1[CL3B] RNA. Lanes 4–6, R1.1[WC-R] RNA. The latter RNA migrates faster in the gel due to a persistent secondary structure. Lanes 7–9, R1.1 RNA. Lanes 3, 6 and 9 represent the products of incubation with TbCl3. Lanes 1, 4 and 7 show an alkaline ladder obtained by heating 5' 32P-labeled RNA at 90°C for 10 min in 1 mM sodium carbonate buffer (pH 9.2). Lanes 2, 5 and 8 represent partial RNase U2 (A>G-specific) reactions, obtained by incubating RNA at 55°C for 15 min with 10 U of RNase U2 in 25 mM sodium citrate (pH 3.5), 5 M urea, 0.75 mM EDTA and 0.5 mg/ml tRNA. Shown are the positions of the R1.1 internal loop (IL) sequences and the tetraloop sequence. (B) Mfold structures for R1.1 RNA and R1.1[CL3B] RNA, also showing the sites of significant cleavage by Tb3+.

 
For R1.1 RNA, the cleavage pattern in the internal loop region is qualitatively distinct from that of the corresponding region of R1.1[CL3B] RNA. In particular, the 5' proximal segment of the internal loop (A17-C20) is uniformly reactive to Tb3+, with significant cleavage also occurring at U45 and U47 in the 3' proximal segment of the internal loop (Fig. 6A, lane 9; summarized in Fig. 6B). The reactivity pattern of R1.1 RNA is consistent with an unstructured internal loop, as also indicated by the Mfold analysis and by the absence of stable hydrogen bonds within the loop (25). For R1.1[WC-R] RNA, the absence of significant Tb3+ cleavage in the region corresponding to the internal loop (Fig. 6A, lane 6) is distinct from the R1.1[CL3B] RNA pattern, and is consistent with the presence of a regular double-helical structure. It should also be noted that for all three RNAs, conservation of an overall hairpin structure is indicated by strongly reactive sites in the tetraloop and flanking nucleotides.

Both bulged nucleotide positions in R1.1[CL3B] RNA are required for uncoupling
The bulge–helix–bulge motif is implicated in uncoupling, as it is the distinguishing feature of R1.1[CL3B] RNA. The three GC bp most likely are not directly responsible for uncoupling, as revealed by the reactivity of an R1.1 RNA variant in which the internal loop is replaced by four GC base pairs (20). It therefore is likely that one or both of the bulges is required to suppress cleavage. Two variants of R1.1[CL3B] RNA were prepared which carried either a deletion of A20 or a deletion of A48 and U49. A third variant was synthesized in which all three nucleotides were deleted. A cleavage assay reveals that deletion of either bulge is sufficient to allow cleavage (Fig. 7, compare lanes 6 and 8 with lane 4). However, the variant exhibiting a deletion of both bulge positions exhibits the greatest extent of cleavage (Fig. 7, lane 10). For all three R1.1[CL3B] RNA variants, the product sizes are consistent with RNase III cleavage of the primary and secondary R1.1 RNA cleavage sites.



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  		Figure 7. Both sets of bulged nucleotides in R1.1[CL3B] RNA are required to uncouple binding and cleavage. RNAs were synthesized in 5' 32P-labeled form (see Materials and Methods). RNAs were incubated in the presence or absence of RNase III (50 nM) for 10 min at 37°C, then electrophoresed in a denaturing polyacrylamide gel and analyzed by phosphorimaging (see Materials and Methods). Lanes 1 and 2, R1.1 RNA. Lanes 3 and 4, R1.1[CL3B] RNA. Lanes 5 and 6, R1.1[CL3B;{Delta}A48] RNA. Lanes 7 and 8, R1.1[CL3B;{Delta}A48{Delta}U49] RNA. Lanes 9 and 10, R1.1[CL3B;{Delta}A20;{Delta}A48{Delta}U49] RNA. Note the significantly greater electrophoretic mobility of the deletion variants, due to their shorter lengths and perhaps also to an altered secondary structure in the presence of 7 M urea.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
This study has described the isolation and characterization of a T7 R1.1 RNA variant that is recognized by RNase III but is resistant to cleavage. An in vitro selection protocol allowed the isolation of cleavage-resistant sequences from a R1.1 RNA pool containing a sequence-randomized internal loop. Gel shift analysis of selected RNAs identified R1.1[CL3B] RNA as a species with a binding affinity comparable to that of R1.1 RNA. Mfold analysis and Tb3+ ion structure probing indicate that R1.1[CL3B] RNA possesses a bulge–helix–bulge motif in place of the unstructured, [4/5] asymmetric internal loop. The restoration of cleavage reactivity by deletion of either bulge demonstrates that both bulges are necessary to confer resistance. The precise structure of the motif remains to be determined. For example, there are two alternatives to the A20 bulge, involving either A21 or A22. It is also not known whether bulge nucleotide identity is important for uncoupling, or whether the bases are extrahelical or intrahelical. R1.1[CL3B] RNA undergoes a minor amount of cleavage (~5%) at high enzyme concentrations and extended reaction times. It is therefore possible that R1.1[CL3B] RNA variants may exhibit an even greater resistance to cleavage and/or stronger binding affinity. The low reactivity prevented an accurate determination of the Km and kcat for RNase III cleavage of R1.1[CL3B] RNA. However, based on the gel shift behavior (this study) and the similar KD and Km values for E.coli RNase III binding and cleavage of R1.1 RNA, respectively (19,40), the inhibition of cleavage is expected to reflect a reduction in kcat.

The mechanism of substrate-dependent uncoupling is unclear. R1.1[CL3B] RNA may exhibit a specific type of ‘dsRNA mimicry’—a term originally applied to describe the sustained reactivity of internal loop-containing substrates towards an otherwise strictly double-strand-specific nuclease (18). The bulge–helix–bulge motif would permit RNase III recognition, but in contrast to a regular double-helical structure (or a [4/5] internal loop), it would block productive engagement of the scissile phosphodiesters at the active site, perhaps by hindering a conformational change. In this regard, a structural change in RNase III has been detected in association with the catalytic step (40). The mechanism of uncoupling ultimately will require a structural analysis of an RNase III–R1.1[CL3B] RNA complex.

Substrate structure-dependent uncoupling of E.coli RNase III suggests that RNase III orthologs in general could function as non-catalytic, RNA-binding proteins. Transplantation of the bulge–helix–bulge motif into heterologous substrates could provide one test for this proposal. Alternatively, structurally distinct motifs may be necessary to uncouple RNase III ortholog action. A cleavage-resistant RNA recognized by RNase III may provide a novel gene-regulatory element. Through recruitment of RNase III as an RNA-binding protein, an mRNA could be stabilized or translationally regulated by a 5'- or 3'-end-localized RNA–protein complex. Expression of a cleavage-resistant, binding-competent RNA may exert a dominant-negative effect on RNase III function in vivo.

RNA processing antideterminants are proposed to play key roles in the site-specificity of ribonuclease action as well as protect functionally important sequences from inadvertent cleavage (9,20,41). Since the bulge–helix–bulge motif of R1.1[CL3B] RNA does not block RNase III recognition, it may be regarded an example of a ‘catalytic’ antideterminant for RNase III. This can be distinguished from recognition antideterminants, which block cleavage by inhibiting RNase III binding (20).

The 5' untranslated leader of the phage lambda cIII mRNA has been proposed to bind RNase III, with this event serving to activate cIII protein production (27). This study supports the feasibility of such a mechanism, and further analysis of the RNase III–cIII mRNA interaction is warranted. An Mfold analysis of the cIII mRNA 5' leader does not reveal a bulge–helix–bulge motif similar to that of R1.1[CL3B] RNA (I.C.-J. and A.W.N., unpublished results). However, other structures may exist which also can uncouple RNase III action.

The cleavage resistance of the Class I R1.1 RNA variants and the involvement of the A9U mutation also are of interest. Mfold analysis of selected Class I R1.1 RNA variants indicates a novel base-pairing interaction involving C residues in the 5' segment of the internal loop and the three consecutive G residues at positions 10–12 (see Fig. 8). This pairing creates a stem–loop structure in place of the R1.1 lower stem and internal loop. Disruption of the internal loop and lower stem would be sufficient to block RNase III recognition and cleavage, and also would explain the altered nondenaturing gel electrophoretic mobilities of the Class I RNAs. Mfold also reveals that the A9U mutation is required for the alternative conformation. The A9U mutation may destabilize the R1.1 lower stem by creating a UU mismatch, thereby allowing the pairing of the C-rich 5' proximal segment of the internal loop with the consecutive G residues. The inability of the A9U mutation on its own to inhibit cleavage may reflect an insufficient number of C residues in the R1.1 internal loop to allow the alternative base-pairing. Further structural analysis of Class I R.1 RNA variants should provide a critical test of the existence of the proposed conformation, and provide further insight on structure–reactivity correlates of RNase III substrates.



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  		Figure 8. Proposed alternative conformation of the Class I variant R1.1[CL68B] RNA, and the involvement of the A9U mutation. On the left is R1.1[CL68B] RNA lacking the A9U mutation. Shown are the positions of the proximal and distal boxes, and the RNase III cleavage site. In the presence of the A9U mutation, the bracketed nucleotides are proposed to pair, producing the structure shown on the right. Note the disruption of the proximal box (pb), as well as loss of the R1.1 internal loop and lower stem.

 

   ACKNOWLEDGEMENTS
 
The authors thank Dr Philip Cunningham for generously allowing the use of his DNA sequencer and sequence analysis software. We also thank Marnie Waddington and Ashesh Suraiya for technical assistance in sequencing. The authors also acknowledge helpful discussions with Dr Nils Walter (University of Michigan) regarding terbium ion structure probing protocols, and Dr Alexandre Pertzev for sharing unpublished information on distal box mutations. The authors also thank other members of the laboratory for their advice and encouragement. This project was supported by NIH grant GM56772.


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