Characterizing the Structural Features of RNA/RNA Interactions of the F-plasmid FinOP Fertility Inhibition System*

Michael J. Gubbins {ddagger} §, David C. Arthur ¶, Alexandru F. Ghetu ¶, J. N. Mark Glover ¶ and Laura S. Frost {ddagger} ||

From the {ddagger}Department of Biological Sciences, University of Alberta, Edmonton, Alberta T6G 2E9, Canada and The Department of Biochemistry, University of Alberta, Edmonton, Alberta T6G 2H7, Canada

Received for publication, March 27, 2003 , and in revised form, May 12, 2003.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
F-like plasmid transfer is mediated by the FinOP fertility inhibition system. Expression of the F positive regulatory protein, TraJ, is controlled by the action of the antisense RNA, FinP, and the RNA-binding protein FinO. FinO binds to and protects FinP from degradation and promotes duplex formation between FinP and traJ mRNA, leading to repression of both traJ expression and conjugative F transfer. FinP antisense RNA secondary structure is composed of two stem-loops separated by a 4-base single-stranded spacer and flanked on each side by single-stranded tails. Here we show that disruption of the expected Watson-Crick base pairing between the loops of FinP stem-loop I and its cognate RNA binding partner, traJ mRNA stem-loop Ic, led to a moderate reduction in the rate of duplex formation in vitro. In vivo, alterations of the anti-ribosome binding site region in the loop of FinP stem-loop I reduced the ability of the mutant FinP to mediate fertility inhibition and to inhibit TraJ expression when expressed in trans at an elevated copy number. Alterations of intermolecular complementarity between the stems of these RNAs reduced the rate of duplex formation. Our results suggest that successful interaction between stem-loop I of FinP and stem-loop Ic of traJ mRNA requires that base pairing must proceed from an initial loop-loop interaction through the top portion of the stems for stable duplex formation to occur.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
RNA/RNA interactions are important in a wide variety of biological systems (reviewed in Refs. 1 and 2). Antisense RNA interactions are key components of plasmid replication control for the Escherichia coli plasmids ColE1 (35), R1 (6, 7), and the paradigm for pilus-mediated plasmid transfer, F (reviewed in Ref. 8). Most of the genes required for conjugative F plasmid transfer are encoded in the 33.3-kb transfer (tra) operon, and expression of the operon is driven from the major tra operon promoter, PY (reviewed in Ref. 8). Efficient transcription from PY requires the activity of the positive regulatory protein, TraJ (9). TraJ expression is in turn regulated by the activity of the antisense RNA, FinP (GenBankTM accession number U01159 [GenBank] ), and the RNA-binding protein, FinO (Protein Data Bank accession number 1DVO [PDB] ). FinP antisense RNA is composed of 79 bases arranged into two stem-loops (SL)1 separated by a 4-base single-stranded spacer and flanked by 4-base and 6-base single-stranded tails on the 5' and 3' sides, respectively (Fig. 1) (10). FinP is complementary to a portion of the 5'-untranslated region (UTR) of traJ mRNA, which contains SL-Ic and SL-IIc, complementary to FinP SL-I and SL-II, respectively (Fig. 1) (1113). The traJ ribosome binding site (RBS) is located in the top portion of SL-Ic (Fig. 1), and binding of FinP to the 5'-UTR of traJ mRNA is believed to sequester the RBS within a FinP/traJ mRNA duplex, via base pairing with a region in the loop of FinP SL-I termed the anti-RBS. This interaction is thought to prevent TraJ translation, thus repressing expression of the F plasmid transfer operon (11, 14, 15).



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FIG. 1.
Secondary structure of FinP antisense RNA and a portion of the 5'-UTR of traJ mRNA. Only a portion of the 5' UTR of traJ that is complementary to FinP antisense RNA is shown for clarity. A full representation of the traJ 5'-UTR is available in Ref. 21. Black lines indicate the traJ RBS and the corresponding anti-RBS of FinP. The traJ start codon is shown in italic type. Nucleotide positions are labeled at every tenth base from the 5'-end of the molecules.

 

In F-like plasmids, the regulatory activity of FinP depends upon the action of the plasmid-encoded protein, FinO (16), and in F, whose finO gene is interrupted by an IS3 insertion element, transfer is completely derepressed (17). FinO is not plasmid-specific, and when supplied in trans, FinO from the related plasmid R6–5 (18) or R100 (19) can repress F transfer. FinO is a 186-amino acid, 21.2-kDa basic cytoplasmic protein with a highly {alpha}-helical structure that adopts a novel protein fold (20). FinO binds the relatively unstable FinP molecule (2022), sterically inhibiting RNase E cleavage of the single-stranded spacer between SL-I and SL-II (23, 24) and allowing the steady-state concentration of FinP to increase to sufficient levels to mediate repression of traJ expression (25, 26). Indeed, the requirement of FinO for inhibition of transfer and traJ expression can be alleviated by providing FinP at elevated copy number in the cell (14, 15). FinO also catalyzes FinP/traJ mRNA duplex formation in vitro (10, 20, 23), which is believed to allow rapid sequestration of the traJ RBS and efficient inhibition of traJ expression in vivo (14, 15).

"Kissing" between loops of RNA stem-loop structures is commonly the first interaction to occur during the process of RNA/RNA duplex formation (reviewed in Refs. 27 and 28). F-like conjugative plasmids encode eight different alleles of FinP, with the highest variability in the loops (8, 12, 21). The loop sequences of FinP and traJ mRNA are therefore thought to be responsible for mediating the plasmid specificity of the F-like FinOP systems (12, 14, 15) and are thought to be the initial site of interaction between the sense and antisense RNAs. Although the loop sequences of FinP in F-like plasmids vary, a common motif, 5'-YUNR-3' (where Y represents pyrimidine, N is any base, and R is purine), is found in several finP alleles (8), which is a key structural motif in the loops of many antisense RNA molecules (29).

The structural features of SL-I and SL-Ic of FinP antisense RNA and traJ mRNA, respectively, that influence FinO-mediated duplex formation in vitro were characterized. Duplex analyses employing EMSAs using in vitro synthesized RNAs and purified FinO protein were used to measure apparent second-order association rate constants (kapp) of duplex formation for a variety of interacting RNA partners. Our studies demonstrate that both in vitro and in vivo, FinO can overcome a variety of sequence and structural changes to FinP SL-I and traJ mRNA SL-Ic in order to promote duplex formation.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Preparation of in Vitro Transcription Templates—All RNAs employed in this work are derived from F-encoded FinP and traJ mRNA. All solutions used for RNA work were treated with diethyl pyrocarbonate prior to use. Transcription of FinP SL-I, traJ SL-Ic, and their mutant derivatives employed the T7 RNA polymerase promoter top strand oligonucleotide primer MGU5 and various oligonucleotide primer templates (Table I). Oligonucleotide template primers (Molecular Biology Services Unit, University of Alberta) were electrophoresed on a 10% (29:1), 8 M urea polyacrylamide gel at a constant 300 V for ~2 h. Full-length oligonucleotides were excised from the polyacrylamide gels after ethidium bromide staining. The gel slices were crushed and eluted overnight at 37 °C in elution buffer (0.5 M NH4OAc, 0.1 mM EDTA). The supernatants were removed and extracted with an equal volume of phenol/chloroform (1:1), followed by an extraction with chloroform. The DNA was then ethanol-precipitated and dissolved in 30–50 µl of Milli-Q water, and the concentration of the purified DNA was determined by UV absorbance at 260 nm in a Bio-Rad Smart-Spec 3000. Seventy-five pmol of the template strand were mixed with a 1.5-fold molar excess of the complementary T7 RNA polymerase promoter top strand oligonucleotide primer, MGU5, in a total volume of 50 µl. MgCl2 was added to a final concentration of 100 mM, and the mixture was heated to 95 °C for 5 min. Cooling to room temperature over 1 h and 45 min allowed annealing of the template oligonucleotide to the MGU5 T7 RNA polymerase primer.


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TABLE I
Primers used in this study

Base changes are underlined, and the T7 promoter sequence is in boldface type.

 

In Vitro Transcriptions—Annealed templates prepared as described above were added to a final concentration of 300 nM in 20-µl transcription reactions. For labeled reactions, GTP/ATP/CTP were added to a final concentration for each of 2.5 mM, and UTP was added to a final concentration of 0.1 mM, along with 10–50 µCi of [{alpha}-32P]UTP (PerkinElmer Life Sciences). Twenty-six units of RNA Guard (Amersham Biosciences) were added to each reaction, along with 1x transcription buffer (Roche Applied Science) supplemented with 0.01% (v/v) Triton X-100 (Sigma) and 0.1 mg/ml bovine serum albumin (RNase Free; Roche Applied Science). Twenty units of T7 RNA polymerase (Roche Applied Science) were added, and the reactions were incubated at 37 °C for 2 h. Ten units of DNase I (RNase Free; Roche Applied Sciences) were added, and the reactions were incubated for a further 15 min to digest the template DNA. One-fifth volume of RNA load dye (96% (v/v) deionized formamide, 0.05% (w/v) each xylene cyanol and bromphenol blue, 20 mM EDTA) was added, and the RNA was heated to 95 °C for approximately 5 min and then cooled on ice. The RNA was electrophoresed on an 8% (29:1), 8 M urea polyacrylamide gel at 250 V for ~2 h. The radioactive RNA band was visualized by exposure to Kodak X-Omat R film for several minutes and then excised and purified as described above, except the purified RNA was dissolved in 10 µl of Milli-Q water after precipitation. To make unlabeled RNA, all procedures were the same, except GTP/ATP/CTP/UTP were added to transcription reactions at a final concentration of 2.5 mM. The unlabeled RNA was visualized by staining in ethidium bromide and gel-purified as described above.

EMSAs for Apparent Second Order Association Rate Constant Determination—EMSA analyses for determination of kapp values were performed essentially as described (20, 23). Briefly, 60 fmol of 32P-labeled RNA was incubated with 600 fmol of its unlabeled complementary RNA in a 50-µl reaction containing 1x TMN buffer (20 mM Tris-HCl, pH 7.5, 10 mM magnesium acetate, 100 mM NaCl). Plasmid R6–5 FinO, purified as described (22), was added to a final concentration of 6 µM to the reactions where appropriate. Reactions were incubated at 37 °C, and 5-µl aliquots were withdrawn at various times, mixed with 10 µl of ice-cold TMN stop solution (1x TMN containing 30% (v/v) glycerol and 0.05% (w/v) bromphenol blue), and kept on ice. The samples were then electrophoresed on 8% (29:1) nondenaturing polyacrylamide gels containing Tris/glycine buffer (25 mM Tris-HCl, pH 8.0–8.3, 190 mM glycine) at a constant 160 V for 65 min at room temperature. Gels were dried and then exposed on Molecular Dynamics Storage Phosphor screens overnight. Free and duplexed RNA species were visualized and quantified using an Amersham Biosciences PhosphorImager 445 SI and ImageQuaNT software. k1 values were derived from log plots of the percentage of free labeled RNA versus time of incubation to determine the time required for 50% of the free labeled RNA to form a duplex. kapp values were then calculated from k1 and the concentration of the RNA species in excess, essentially as described (23, 30).

EMSA for Detection of FinO Binding to SL-I and SL-I({Delta}tails)—The association equilibrium constant (Ka) for FinO binding to 32P-labeled FinP SL-I or derivatives thereof was performed as described (22), except 6 fmol of 32P-labeled RNA were used per reaction instead of 7.5 fmol. Quantification of unbound and FinO-bound RNAs and calculation of the association constants were performed exactly as described (22, 23).

Construction of Recombinant Plasmids—The plasmids used in this study and the relevant details and sources of each are listed in Table II. Isolation of all plasmid DNA was performed using a rapid alkaline extraction technique (31). All clones constructed during the course of this work were sequenced using the DYEnamic ET fluorescent sequencing system according to the manufacturer's instructions (Amersham Biosciences) to confirm that the correct DNA sequence was present in each clone. All restriction enzymes used for DNA cloning were purchased from Roche Applied Science. The plasmid pUC180GGA was constructed to express FinP(C16G/C17G/U18A) from its own promoter from the high copy number vector pUC18. The plasmid pUC180 contains a 180-base EcoRI/HindIII fragment derived from F, which encodes wild-type FinP, expressed from its own promoter in the absence of transcription from PtraJ (24). This plasmid served as a template for site-directed mutagenesis of FinP using the mutagenic primers MGU53 and MGU54 (Table I) and Pfu Turbo® (Stratagene) to create pUC180GGA. All procedures were performed according to the manufacturer's instructions (Stratagene), except the plasmid was transformed into rubidium chloride-competent E. coli to propagate the DNA (32). The presence of the mutation was confirmed by sequencing as described above. pLT180GGA was created by inserting the 180-base EcoRI/HindIII fragment containing FinP(C16G/C17G/U18A) from pUC180GGA into similarly digested pT7-3 (33), allowing this mutant FinP antisense RNA to be expressed from its own promoter in a moderate copy number plasmid.


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TABLE II
Plasmids and bacterial strains used in this study

 

Mating Assays—E. coli MC4100 containing the finP F-derivative plasmid pSLF20 (Table II) was transformed with the control plasmid pT7-3 or one of the test plasmids (pLT180, pLT180GGA) that express FinP in trans from its own promoter (Table II). To provide FinO in trans, the plasmid pSnO104 (Table II) was transformed into the test strains where appropriate. Donor cultures were grown with appropriate antibiotic selection at 37 °C with agitation to midlog phase (A600 ~0.6–1.0), and 0.5 A600 equivalents were pelleted and washed with fresh LB broth (Difco) to remove antibiotics and then resuspended in 500 µl of fresh LB broth. The recipient strain ED24 (Table II) was grown to midlog phase without antibiotic selection. Aliquots (100 µl) of donor and recipient cultures were mixed with 800 µl of fresh LB and then incubated at 37 °C for 45 min with no agitation. Cultures were vortexed vigorously for 15 s and placed on ice to disrupt mating pairs. 10-fold serial dilutions were made using ice-cold 1x PBS (137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4·7H2O, 1.4 mM KH2PO4), and 10 µl of each dilution were inoculated on agar plates to select for donor and transconjugant cells. Donors containing pSLF20 alone were selected on Maconkey-Lactose plates (Difco) containing 200 µg/ml streptomycin. Donors containing pSLF20 and any one of the plasmids pT7-3, pLT180, or pLT180GGA were selected on Maconkey-Lactose plates containing 200 µg/ml streptomycin and 5.0 µg/ml ampicillin. All donor constructs containing pSnO104 were selected on Maconkey-Lactose plates to which chloramphenicol was added to a final concentration of 50 µg/ml, in addition to the other antibiotics listed above. All transconjugants were selected on L1-spectinomycin plates (25) containing 100 µg/ml spectinomycin. All plates were incubated at 37 °C for 12–36 h until visible colonies appeared. Donor and transconjugant cells were then counted, and the ratio of transconjugants to donors was calculated, allowing mating efficiency to be compared with the control of conjugative transfer of pSLF20 alone, which was set at 100% mating efficiency.

Immunoblot Analysis—Cell pellets corresponding to 0.1 A600 equivalents were boiled in SDS sample buffer (34) for 5 min, and the supernatants were electrophoresed on 15% (29:1) SDS-polyacrylamide gels using the Bio-Rad Mini-Protean® system. Proteins were transferred to Immobilon-P membranes (Millipore Corp.) at 100 V for1hat4 °C using Towbin buffer (35). Membranes were blocked overnight at 4 °C with 10% (w/v) skim milk (Difco) dissolved in TBST (50 mM Tris-HCl, pH 7.5, 150 mM NaCl, and 0.1% (v/v) Tween 20 (Caledon Laboratories)). Primary antibodies diluted in 10% skim milk in TBST were added to blots and incubated for 1 h at room temperature. The following dilutions of polyclonal antisera (raised in rabbits) were used: anti-FinO, 1:50,000; anti-TraJ, 1:15,000. Blots were washed at room temperature four times for 15 min each with TBST. The secondary antibody used was horseradish peroxidase-conjugated donkey anti-rabbit IgG (Amersham Biosciences) at a 1:10,000 dilution. Blots were incubated for 1 h at room temperature with the secondary antibody and then washed as described above. Blots were developed with Renaissance Western blot Chemiluminescence Reagent Plus (PerkinElmer Life Sciences) and exposed to Eastman Kodak Co. X-Omat R film for varying times to visualize the signals.

Northern Blot Analysis—Total RNA was isolated via a modified hot phenol method as described (23, 24) from strains grown in liquid cultures (LB broth) at 37 °C to an A600 of 0.8–1.0. RNA (30 µg) was denatured for 5 min at 95 °C in formamide RNA load dye and then electrophoresed on an 8% (29:1), 8 M urea polyacrylamide gel and transferred to Zeta-Probe nylon membranes (Bio-Rad) as described (24). The blots were prehybridized for 4 h using the same conditions as described (24), except 200 µg/ml each of boiled E. coli strain W tRNA type XX and sonicated calf thymus DNA (Sigma) were added to the hybridization solution. The FinP-specific probe primer A (Table I) was 5'-end-labeled with T4 polynucleotide kinase (Roche Applied Science) and [{gamma}-32P]ATP (PerkinElmer Life Sciences), and ~10 pmol of the probe was added to the blots in fresh hybridization solution. Incubation proceeded overnight at 37 °C, and the blots were then washed as described (23) and exposed on an Amersham Biosciences storage phosphor screen. Bands corresponding to FinP were visualized using an Amersham Biosciences PhosphorImager 445 SI and ImageQuaNT software.

RNA Secondary Structure Predictions—Secondary structure predictions and {Delta}G values of SL-I, SL-Ic, and their derivatives were performed using the Mfold version 3.1 algorithm (36, 37). The RNA sequences were analyzed at the Rensselaer Polytechnic Institute Mfold server (available on the World Wide Web at bioinfo.math.rpi.edu/ ~mfold) using standard settings. The secondary structure of FinP and traJ184 mRNA were experimentally determined previously (10) and were used as a reference with which to compare the predicted structures of the individual stem-loop constructs employed in this study.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
FinO from the related F-like plasmid R6-5 has been determined to function in vivo to repress F transfer and in vitro to both bind FinP antisense RNA and promote duplex formation between FinP and traJ mRNA (10, 20, 22). In the absence of FinO, FinP/traJ184 mRNA duplex formation occurs in vitro with a kapp of 5 x 105 M–1 s1, and in the presence of wild-type R6–5 FinO, the kapp increases to 2.5 x 107 M–1 s1 (20). In the present study, a variety of RNA stem-loop constructs derived from FinP and traJ184 mRNA were synthesized in vitro and subjected to EMSA analysis to determine their apparent second order association rate constants in the presence and absence of FinO. In order to determine specific regions of FinP and traJ184 mRNA that are required for duplex formation, multiple sequence and structural mutants of FinP SL-I were synthesized by in vitro T7 RNA polymerase transcription. SL-I was chosen for several reasons. The anti-RBS for traJ mRNA is located in SL-I (Fig. 1), which has been hypothesized to make important initial contacts with the traJ RBS in order to prevent translation of traJ in vivo (15). The loop of SL-I is also slightly larger than the loop of SL-II, and whereas both contain the consensus 5'-YUNR-3' motif, which has been shown to make important structural contributions to RNA/RNA interactions (29), SL-I exhibits higher conservation in the loop nucleotides than SL-II, among the known alleles of FinP (8, 12, 21). These observations suggest that the more highly conserved loop of SL-I is important in the initial interaction between FinP and traJ mRNA molecules and in the inhibition of traJ mRNA translation.

A single base pair mismatch in the stem of SL-I and two single base pair mismatches in the stem of SL-Ic results in lower stability of these stems compared with the more extensively base-paired stems in SL-II and SL-IIc (Fig. 1). The lower predicted free energy of unfolding of SL-I ({Delta}G = –10.1 kcal/mol) and SL-Ic ({Delta}G = –8.6 kcal/mol) compared with SL-II ({Delta}G = –28.2 kcal/mol) and SL-IIc ({Delta}G = –23.3 kcal/mol) suggests that intermolecular base pairing between the stems of SL-I and SL-Ic during the formation of a stable FinP/traJ184 mRNA duplex are more likely to occur than between the stems of SL-II and SL-IIc (10).

Contribution of the Loop Residues of SL-I to RNA/RNA Duplex Formation—Three regions of the loop of SL-I were chosen to test for their contribution to SL-I/SL-Ic duplex formation. All mutations were transversions that disrupted the expected Watson-Crick base pair interactions between the loops. The predicted secondary structures of all of these constructs are shown in Fig. 2. One- and two-base transversion mutations in these regions resulted in no noticeably obvious alterations to duplex formation ability (data not shown); therefore, 3- and 4-base transversion mutations were examined. The first mutation examined lies within the 5' side of the loop of SL-I, 5'-C16G/C17G/U18A-3', which is referred to as SL-I (1618) throughout this work. This mutation alters 3 of the 6 bases that comprise the predicted anti-RBS of FinP (Fig. 1). The second mutation is located on the 3' side of the loop of SL-I, 5'-C21G/A22U/A23U-3', which is referred to as SL-I(21–23). The last extends across the top of the loop of SL-I, 5'-U18A/C19G/A20U/C21G-3', which is referred to as SL-I(18–21). When compared with the kapp for SL-I/SL-Ic duplex formation under identical conditions, the kapp for SL-I(16–18)/SL-Ic duplex formation was reduced by ~52% in the absence of FinO and by 55% in the presence of FinO (Fig. 3, A and B; Table III). SL-I(18–21)/SL-Ic duplex formation exhibited a kapp reduced by 35% in the absence of FinO and by 60% in the presence of FinO, whereas SL-I(21–23)/SL-Ic revealed a kapp reduced by 55% in the absence of FinO and by 51% in the presence of FinO, when compared with the kapp for SL-I/SL-Ic duplex formation under the same conditions (Fig. 3, A and B; Table III). These results suggest that the level of complementarity between loop residues of SL-I and SL-Ic affects FinO-mediated duplex formation in vitro. The observation that the kapp values for duplex formation of all of the interactions tested between the SL-I loop mutants and SL-Ic were 10–19-fold higher in the presence of FinO than in the absence of FinO (Table III) reveals that FinO can overcome as many as four mismatches in the loop-loop base pairing interaction to promote duplex formation in vitro.



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FIG. 2.
In vitro transcribed SL-I and SL-Ic constructs employed in this work. All RNAs were transcribed and purified as outlined under "Experimental Procedures," using the DNA oligonucleotide template primers listed in Table I. Loop and single-base stem mutations are indicated in boldface type. The traJ mRNA start codon in the SL-Ic constructs is indicated in italic type. The regions of the SL-Ic stem that were changed to alter complementarity with the corresponding regions in SL-I are outlined by black boxes. An extra Gly residue at the 5'-end of each molecule, resulting from in vitro transcription, is omitted simply for clarity.

 


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FIG. 3.
Mutations in SL-I that disrupt Watson-Crick base pairing interactions between the loops decrease SL-I/SL-Ic duplex formation rates. A, EMSA analysis of duplex formation between 32P-labeled wild-type SL-I and unlabeled SL-Ic. Samples were taken at 0, 15, 30, 60, and 120 min (–FinO) and at 0, 1, 2, 3, 4, 6, 10, and 15 min (+FinO) after the initiation of duplex formation. Open arrows indicate free RNA, and closed arrows indicate RNA duplexes. Conditions of EMSA analysis and determination of kapp values for duplex formation are described under "Experimental Procedures." B, EMSA analysis of SL-I loop mutants duplexing with SL-Ic. The RNA species present in each reaction are indicated above each panel. In each RNA/RNA pair, the SL-I loop mutant derivatives were 32P-labeled, whereas SL-Ic was unlabeled. EMSA procedures and kapp determination procedures are described under "Experimental Procedures."

 

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TABLE III
kapp values for duplex formation between SL-I and SL-Ic derivatives

 

The Effect of Stem Mutations on SL-I and SL-Ic Duplex Formation—The bulged A12:A27 base pair mismatch in SL-I and the corresponding U85:U100 mismatch in SL-Ic (Fig. 2) were examined for their contribution to duplex formation. SL-I(A27U) ({Delta}G = –14.3 kcal/mol) and SL-Ic(U85A) ({Delta}G = –12.1 kcal/mol) were made to increase the stability of the stems while maintaining full intermolecular complementarity between the stems of the two RNAs. The kapp for SL-I(A27U)/SL-Ic(U85A) duplex formation in the absence of FinO was reduced by 32%, whereas in the presence of FinO, the kapp showed a 74% reduction, compared with the kapp for SL-I/SL-Ic duplex formation (Fig. 4A, Table III). These results suggest that the overall stability of the stem regions of SL-I and SL-Ic influences their transition to a stable duplex. To create more drastic mutations affecting stem complementarity and to provide insight into the direction of progression of duplex formation, SL-Ic(TSR) and SL-Ic(BSR) were constructed. SL-Ic(TSR) has 5 base pairs in the stem immediately below the loop reversed in orientation, resulting in noncomplementarity with the corresponding region in SL-I (Fig. 2). SL-Ic(BSR) has 6 base pairs at the bottom of the stem reversed in sequence in the same fashion (Fig. 2). The single-stranded tail regions were not included in these constructs, ensuring that only the effects on intermolecular stem/stem interactions were examined. SL-I/SL-Ic(TSR) duplex formation in both the presence and absence of FinO was minimal, and a kapp could not be calculated in either case because less than 20% of the 32P-labeled free RNA in the reactions was converted to a duplex (Fig. 4A). SL-I/SL-Ic(BSR) duplex formation in the absence of FinO revealed a kapp that was reduced by 84% relative to the kapp for SL-I/SL-Ic duplex formation (Fig. 4A; Table III). In the presence of FinO, the kapp for SL-I/SL-Ic(BSR) duplex formation was reduced by 66% compared with the kapp for SL-I/SL-Ic duplex formation (Fig. 4A; Table III). These results suggest that stable duplex formation between SL-I and SL-Ic can proceed only if intermolecular complementarity extends from the loop through the top of the stem. The virtually identical kapp values for SL-I/SL-Ic({Delta}tails) and SL-I/SL-Ic(BSR) duplex formation also suggests that a region of noncomplementarity at the bottom of the stem has no significant effect on the ability of FinO to promote duplex formation between these constructs in vitro.



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FIG. 4.
Mutations in the stem regions of SL-I and SL-Ic decrease duplex formation rates. A, EMSA analysis of duplex formation between 32P-labeled SL-I or SL-I(A27U) and their unlabeled SL-Ic-derived cognate binding partners (indicated above each panel). In all reactions without FinO (–FinO), samples were taken at 0, 15, 30, 60, and 120 min. In reactions containing FinO (+FinO), samples were taken at 0, 1, 2, 3, 4, 6, 10, and 15 min (SL-I(A27U)/SL-Ic(U85A)) and at 0, 1, 2, 5, 10, 15, 30, and 60 min (SL-I/SL-Ic(TSR) and SL-I/SL-Ic(BSR)). Procedures for EMSA analysis and kapp determination are described under "Experimental Procedures." The open arrows indicate free RNA, and closed arrows indicate RNA duplexes. B, EMSA analysis to detect a kissing intermediate between 32P-labeled SL-I and unlabeled SL-IcR. Samples were taken at 0, 60, and 120 min, in both the presence and absence of FinO, as indicated above each panel, and analyzed via EMSA analysis as described under "Experimental Procedures." The open arrow indicates free RNA.

 

Detection of SL-I/SL-Ic Kissing Complexes—Since kissing between loop regions is normally the first interaction in most antisense/sense RNA-pairing reactions, it was decided to determine whether a SL-I/SL-Ic kissing dimer could form and be detected by EMSA analysis. SL-IcR was created such that the loop region was completely complementary to SL-I, but the stems and tails were not complementary (Fig. 2). In the presence and absence of FinO, no stable kissing intermediate was detectable (Fig. 4B), suggesting that that any initial kissing complex that forms between SL-I and SL-Ic is transient and unstable and is not detectable by EMSA analysis. These results also confirm the observations resulting from the SL-I/SL-Ic-(TSR) duplexing experiments described above. The formation of a stable SL-I/SL-Ic duplex requires complementarity in both the loops and as much as half of the stem in the region immediately below the loops of both RNA molecules.

Contribution of the Single-stranded Tail Regions of SL-I to RNA/RNA Duplex Formation—Since the single-stranded tails of FinP SL-I and SL-II have been shown to influence the ability of FinO to bind FinP with high affinity (21), the contribution of these regions to duplex formation in vitro was tested. SL-I/SL-Ic({Delta}tails) duplex formation showed a kapp reduced by 68% in the absence of FinO, and a kapp reduced by 72% in the presence of FinO, relative to the kapp for SL-I/SL-Ic duplex formation under identical conditions (Fig. 5A; Table III). Analysis of SL-I({Delta}tails) duplex formation with SL-Ic({Delta}tails) revealed a 55% decrease in kapp in the absence of FinO and an 81% reduction in kapp in the presence of FinO, compared with the kapp for SL-I/SL-Ic duplex formation under identical conditions (Fig. 5A; Table III). These values are comparable with the values obtained for SL-I/SL-Ic({Delta}tails) duplex formation, suggesting that complementarity of the single-stranded tail regions, rather than possible structural alterations to the molecules upon removal of these regions, affects duplex formation in vitro. These results suggest that the presence of the single-stranded regions flanking SL-I and SL-Ic makes important contributions to the FinO-mediated formation of the RNA/RNA duplex in vitro. In order to ensure that any decrease in kapp was the result of alterations in complementarity of the interacting RNAs and not due to an inability of FinO to bind them, EMSA analysis was performed to determine whether FinO could bind to SL-I and SL-I({Delta}tails). As shown in Fig. 5B, FinO was able to bind to both RNA molecules, with a Ka of ~8.6 x 106 M–1 and 3.5x106 M–1, for binding SL-I and SL-I({Delta}tails), respectively. These Ka values are higher than those reported in a previous study, which may be attributable to the fact that our study employed native FinO, whereas the previous study employed a glutathione S-transferase-FinO fusion protein (21). Regardless, our results confirm that FinO could bind the SL-I constructs employed in the duplex assays.



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FIG. 5.
Removal of the single-stranded tails of SL-I and SL-Ic reduces the rate of duplex formation. A, EMSA analysis of duplex formation between 32P-labeled SL-I or SL-I({Delta}tails), and unlabeled SL-Ic({Delta}tails) as indicated above each panel. Samples were taken at 0, 15, 30, 60, and 120 min (–FinO) and at 0, 1, 2, 3, 4, 6, 10, and 15 min (+FinO) after the initiation of duplex formation. EMSA conditions and the procedure for kapp determination are described in detail under "Experimental Procedures." The open arrows indicate free RNA, and closed arrows indicate RNA duplexes. B, EMSA to detect the ability of FinO to bind SL-I and SL-I({Delta}tails). 32P-Labeled RNA (listed above each panel) was incubated with increasing concentrations of FinO (0, 0.038, 0.076, 0.19, 0.38, 0.76, and 1.9 µM) and analyzed by EMSA analysis. The open arrows indicate free RNA, whereas closed arrows indicate FinO-RNA complexes. Procedures for EMSA analysis and determination of the Ka for FinO binding are described under "Experimental Procedures."

 

Contribution of the Anti-RBS of FinP to Its in Vivo Function—As described above, alteration of a portion of the anti-RBS of FinP within the loop of SL-I moderately reduced the efficiency of duplex formation with its complementary RNA, SL-Ic, in vitro. Previous studies have shown that the loop nucleotides of SL-I and SL-II of plasmid R1 FinP directly affect the ability of FinP to inhibit plasmid transfer (14, 15). We wanted to determine the effect of mutations in the anti-RBS of FinP on inhibition of TraJ expression and F plasmid transfer. The plasmid pLT180GGA (Table II) expresses FinP (1618) from a moderate copy number plasmid (pT7-3; 10–30 copies/cell). This plasmid was introduced into E. coli MC4100, with or without plasmid pSnO104, which expresses plasmid R6–5 FinO in trans (Table II). Plasmid pLT180, which expresses wild-type FinP at a moderate copy number, was tested in the same way, as was the negative control parental plasmid, pT7-3 (Table II). The finP F-derivative pSLF20 (Table II) (25) was present in all strains, and pSLF20 conjugative transfer and expression of TraJ were tested in the presence of the FinP derivatives as described under "Experimental Procedures." Mating inhibition assays revealed that FinP (1618), when supplied in trans at medium copy number, was unable to fully repress mating (Table IV). Its efficiency was reduced by ~100-fold and 175-fold, in the absence or presence of FinO, respectively, compared with wild-type FinP under the same conditions. The presence of FinO significantly enhanced mating repression mediated by both wild-type FinP and FinP (1618) expressed in trans at medium copy number (Table IV). Wild-type FinP (pLT180) fully repressed TraJ accumulation in both the presence and absence of FinO, whereas FinP (1618) (pLT180GGA) was able to fully repress TraJ accumulation only in the presence of FinO, as determined by an immunoblot analysis (Fig. 6). These results confirm previous observations that the ability of FinP to inhibit conjugative transfer of F-like plasmids is dependent upon gene dosage (15). They also confirm the results from the in vitro duplex formation assays described above, which showed that FinO can overcome multiple base mutations in FinP SL-I and promote SL-I/SL-Ic duplex formation in vitro when complete loop-loop complementarity is absent. In vivo, it also appears that FinO can compensate for suboptimal loop-loop base complementarity and promote fertility inhibition when the tested FinP loop mutant is supplied at an elevated copy number. Experiments with the very high copy number construct pUC180 gave similar results as with pLT180 (data not shown).


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TABLE IV
Inhibition of pSLF20 conjugative transfer by FinP expressed in trans from medium copy number plasmids

 


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FIG. 6.
Mutation of the anti-RBS of FinP alters its regulatory function in vivo. Shown is immunoblot analysis of TraJ expressed from pSLF20 in the presence and absence of wild-type FinP and FinP (16–18) expressed in trans at moderate copy number. The finP F-derivative plasmid pSLF20 is present in all strains (+) except the negative control lane containing MC4100, which contains no plasmids (–). The presence (+) or absence (–) of pSnO104, which expresses plasmid R6–5 FinO in trans, is indicated above each lane. The presence of the control plasmid pT7-3 or the FinP-expressing plasmids, pLT180 and pLT180GGA, is indicated above the relevant lanes. The location of TraJ and FinO are indicated to the right, and relevant molecular mass protein markers (kDa) are listed on the left. The bottom panel is a Northern blot of total cellular RNA isolated from the same strains, probed with a FinP-specific probe. The location of FinP is indicated on the right. Procedures for immunoblotting and Northern blot analysis are described in detail under "Experimental Procedures."

 


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
This report describes the structural features of FinP antisense RNA and traJ mRNA that influence FinO-mediated duplex formation. A common theme among antisense-sense pairing is the important initial interaction between single-stranded complementary loops of the RNA molecules (4, 30, 38, 39) (reviewed in Ref. 28). For example, in the case of plasmid ColIb-P9, multiple single-base mutations that altered canonical loop-loop base pairing between Inc RNA and RepZ mRNA significantly decreased their in vivo function and impaired RNA/RNA duplex formation in vitro (40). Similarly, a single base mutation that interrupted expected loop-loop pairing between plasmid R1 CopA antisense RNA and its target, CopT, inhibited the formation of a duplex in vitro (30). Three regions of the loop of FinP SL-I were examined for their contribution to FinP/traJ mRNA duplex formation in vitro. Three-base transversion mutations that disrupted expected Watson-Crick base pairing were made in FinP SL-I on the 5' and 3' sides of the loop, whereas a 4-base transversion was made across the top of the loop. In each case, the kapp for duplex formation decreased by a moderate amount, in both the presence and absence of FinO (Table III). This observation suggests that other factors, separate from loop-loop interactions between FinP and traJ mRNA, affect duplex formation.

The plasmid R1 encodes a FinOP system very similar to the F plasmid. Single-base mutations in the top portions of the loops of R1 FinP SL-I or SL-II that altered potential loop-loop base interactions with traJ mRNA significantly inhibited FinO-mediated repression of conjugative transfer of R1, when these mutant FinP molecules were supplied in trans at elevated copy number. However, FinO was able to mediate repression of traJ expression under the same conditions, as measured by {beta}-galactosidase assays of a traJ-lacZ translational fusion reporter construct (15). Whereas mutations in FinP can severely inhibit R1 fertility inhibition, FinO appears to be able to promote direct inhibition of translation of traJ by mutant FinP RNA. When single base transversions were made in the loops of SL-I and SL-II simultaneously, FinO-mediated repression of both traJ expression and conjugative plasmid transfer were significantly reduced, suggesting that both loops play a role in FinP/traJ mRNA duplex formation (15). Interestingly, a single-base transversion mutation made in the 3' portion of the loop of FinP SL-I had no negative effect on FinO-mediated inhibition of traJ expression or plasmid transfer (15). These results suggest that the interaction of FinP and traJ mRNA in vivo relies more on the bases located at the top of the loops than those situated on the 3' side (15). In the present study, the inhibitory function of FinP in vivo was shown to rely on interactions between the anti-RBS of FinP and the RBS of traJ mRNA. When supplied in trans at medium copy number, FinP (1618) exhibited full negative regulatory function only in the presence of FinO. These observations support the finding that FinO can compensate for loop mutations in its RNA targets and promote duplex formation in vitro and confirm that loop-loop base pairing between the anti-RBS of FinP and the RBS of traJ mRNA is critical for the regulatory function of the FinOP system in vivo. However, it must be stressed that under normal physiological conditions (i.e. wild-type levels of FinP and traJ mRNA), it is unlikely that pairing of such mutant FinP molecules with traJ mRNA would occur because of the relatively low levels of these molecules in vivo. In the present study, the inhibitory effect of mutant FinP on plasmid transfer and traJ expression in vivo probably depends completely on the presence of elevated FinP levels in the cell. Indeed, sequence differences in the loops of FinP are responsible for conferring plasmid specificity to the FinOP system of F-like plasmids (12, 15, 41).

A common structural motif in prokaryotic antisense RNA systems is the 5'-YUNR-3' loop motif, which is thought to provide optimal alignment of bases on the 3' side of the loop with those in a complementary RNA (29). Mutations in the YUNR motif of hok RNA of the plasmid R1 hok/sok postsegregational killing system greatly reduced Sok antisense RNA/hok mRNA duplex formation in vitro, although complementarity between the interacting RNAs was maintained (29). In the present study, two of the three multiple loop mutations in FinP SL-I performed in this work altered the YUNR motif and significantly disrupted complementary Watson-Crick base-pairing interactions but led to only moderate decreases in duplex formation rates (Table III). Whether or not the loop mutations disrupted the YUNR motif, the decrease in in vitro duplex formation rates was approximately equivalent. In all cases, the presence of FinO resulted in higher kapp values for duplex formation, demonstrating its ability to promote duplex formation in vitro between RNAs with suboptimal complementarity in loop regions. These results suggest that whereas loop-loop pairing between FinP and traJ mRNA is important, the sequence, and possibly the structure, of the YUNR motif in the loops may play a smaller role than in other systems.

The presence of short single-stranded tails flanking both the 5' and 3' sides of SL-I influenced the ability of SL-I to duplex with SL-Ic in vitro. The removal of both single-stranded tails from SL-I and SL-Ic led to a decrease in FinO-catalyzed duplex formation, which was more significant than any of the loop mutations that were tested, suggesting that single-stranded regions in FinP and traJ mRNA are critical for efficient duplex formation. However, a reduced affinity between FinO and these RNA constructs cannot be ruled out as having an effect on duplex formation at this time. RNA I/RNA II interaction in ColE1 replication control (4) as well as the CopA/CopT interaction of plasmid R1 (38, 42, 43) rely on interactions between single-stranded regions for full activity, demonstrating the importance of such regions to antisense-sense RNA pairing. Considering the short length of complementary single-stranded regions in FinP and traJ mRNA and the importance of such regions to stable duplex formation in other systems (38, 42, 43), the requirement for complementarity in both the loop and single-stranded tail regions of these RNAs is not unexpected.

The presence of bulged nucleotides and mismatched bases in the stems of interacting RNAs is critical for antisense/sense RNA interactions both in vitro and in vivo for several plasmids. These regions are thought to allow breathing of the stems immediately below the loops in order to allow for efficient progression of stable duplex formation (4446). When the purine:purine mismatches in SL-I and SL-Ic were altered to A:U base pairs, maintaining their complementarity but increasing the predicted free energy of unfolding of the stems, the kapp for FinO-mediated duplex formation decreased relative to duplex formation between wild-type SL-I and SL-Ic. These results indicate that the bulged mismatched base pairs in the stems of SL-I and SL-Ic influence the progression of duplex formation. Alteration of complementarity between the stems of SL-I and SL-Ic revealed that, provided at least 5 base pairs immediately below the loops are complementary, stable duplex formation could occur, albeit at a reduced rate. Interaction of the antisense regulatory RNA DsrA with one of its targets, rpoS mRNA, exhibits a similar requirement. Complementarity between bases in the top of the stem of SL-I in DsrA and a specific region of the upstream leader of rpoS mRNA is required for efficient intermolecular pairing in order to promote translation of rpoS (47). Kissing intermediates formed by interacting RNA stem-loop constructs can often be detected readily by EMSA analysis (48, 49). Our inability to detect a SL-I/SL-IcR kissing intermediate by EMSA analysis under the conditions tested suggests that such a complex may be unstable and short-lived, unless initial loop-loop pairing can progress through the stems to form a more stable duplexing intermediate. Alternatively, a stable kissing intermediate may form but might only be detectable using more sensitive means, such as NMR analysis (50). One cannot exclude the possibility that a stable kissing intermediate, mediated by SL-I/SL-Ic and SL-II/SL-IIc interactions between whole FinP and traJ RNAs, may occur, although this possibility remains to be tested.

Several biological systems employ an accessory protein to promote RNA duplex formation, each using a different mechanism. The Rom protein of ColE1 binds to and stabilizes an initial RNA complex between RNAI and RNAII, driving the reaction toward stable duplex formation (5, 51). The E. coli Hfq protein is thought to form a nucleoprotein complex with Spot42 antisense RNA and its target, galK mRNA, cooperatively facilitating RNA/RNA pairing (52). The NCp7 nucleocapsid protein of HIV-1 has been shown to facilitate dimerization between the stem-loops of the dimerization initiation site of the HIV-1 genomic RNA by converting an initial unstable RNA loop-loop complex to a stable dimer (48, 49, 53). More recently, NCp7 was also shown to transiently melt the secondary structure of portions of the stems of human immunodeficiency virus TAR RNA and its DNA complement, cTAR (54). Clearly, accessory proteins that mediate RNA/RNA interactions use a variety of mechanisms to promote RNA pairing. Based upon its similarities to such systems, previous work done on the FinOP system, and the results presented in this work, we present a preliminary model of the mechanism of FinO-mediated duplex formation. FinO is able to destabilize double-stranded RNA, which, along with its RNA-RNA duplex catalysis activity, has been localized to a lysine-rich region within the N-terminal 44 amino acids of the protein.2 The highest affinity binding sites of FinO are SL-II of FinP and SL-IIc of traJ mRNA, although SL-I is also a target for binding (Fig. 5B) (21). Initial binding of FinP and traJ mRNA by FinO allows their loops to come into close proximity and begin base pairing, whereas its RNA destabilization activity begins to open the stems of both SL-I and SL-II. This destabilization of the stems should alleviate the topological restraints inherent in such RNA/RNA interactions, which impose a kinetic barrier to extended duplex formation (reviewed in Ref. 28). Thus, more extensive intermolecular interactions between FinP and traJ mRNA should occur in the presence of FinO. It is likely that destabilization of SL-II and SL-IIc by FinO is more critical than for SL-I and SL-Ic, considering the lower thermal stability of SL-I and SL-Ic imposed by the presence of mismatches in both of their stems. Once duplex formation initiates at the loops and tops of the stems, the single-stranded tail regions of both RNAs may begin to base pair, leading to extended duplex formation. Alternatively, once it has bound to each of its RNA targets, FinO might induce extended regions of single-stranded RNA via its destabilization activity. This function might be similar to that of the E. coli Hfq RNA chaperone protein, which binds to and partially destabilizes the secondary structure of stem-loops in the small, untranslated regulatory RNA OxyS. This destabilization has been hypothesized to facilitate interaction of OxyS with one of its targets, fhlA mRNA (55). Once a critical level of single-stranded RNA is achieved, rapid formation of a stable duplex between the FinP and traJ mRNA can occur.

An interesting observation that emerges from this work is that in vivo, minor changes to the RNA components of the FinOP system can cause significant changes to its function, although in vitro, structural changes to the portions of the RNAs are tolerated during duplex formation. The interaction between FinP, FinO, and traJ mRNA occurs concurrently with transcription of the tra operon, which is in turn influenced by a variety of other factors, including the concentration of these molecules (8). Likewise, the ability of FinOP to inhibit plasmid transfer does not perfectly correlate with the ability of the system to prevent traJ expression (15). It is likely that a delicate balance of factors influences the ability of FinO to promote the formation of a FinP/traJ mRNA duplex in vivo that ultimately results in inhibition of transcription of the tra operon. The exact mechanism underlying FinO-mediated FinP/traJ mRNA duplex formation remains to be elucidated.


    FOOTNOTES
 
* This work was supported in part by grants from the Canadian Institutes of Health Research and the Alberta Heritage Foundation for Medical Research. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ Supported by studentships from the Canadian Institutes of Health Research and the Alberta Heritage Foundation for Medical Research. Back

|| To whom correspondence should be addressed: Dept. of Biological Sciences, CW-405 Biological Sciences Bldg., University of Alberta, Edmonton, Alberta T6G 2E9, Canada. Tel.: 780-492-0672; Fax: 780-492-9234; E-mail: laura.frost{at}ualberta.ca.

1 The abbreviations used are: SL, stem-loops; UTR, untranslated region; EMSA, electrophoretic mobility shift assay; RBS, ribosome binding site. Back

2 A. F. Ghetu, D. C. Arthur, M. J. Gubbins, R. A. Edwards, L. S. Frost, and J. N. M. Glover, submitted for publication. Back



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