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J Bacteriol. 2011 April; 193(8): 1953–1965.
PMCID: PMC3133033

Identification of a Chitin-Induced Small RNA That Regulates Translation of the tfoX Gene, Encoding a Positive Regulator of Natural Competence in Vibrio cholerae[down-pointing small open triangle]


The tfoX (also called sxy) gene product is the central regulator of DNA uptake in the naturally competent bacteria Haemophilus influenzae and Vibrio cholerae. However, the mechanisms regulating tfoX gene expression in both organisms are poorly understood. Our previous studies revealed that in V. cholerae, chitin disaccharide (GlcNAc)2 is needed to activate the transcription and translation of V. cholerae tfoX (tfoXVC) to induce natural competence. In this study, we screened a multicopy library of V. cholerae DNA fragments necessary for translational regulation of tfoXVC. A clone carrying the VC2078-VC2079 intergenic region, designated tfoR, increased the expression of a tfoXVC::lacZ translational fusion constructed in Escherichia coli. Using a tfoXVC::lacZ reporter system in V. cholerae, we confirmed that tfoR positively regulated tfoXVC expression at the translational level. Deletion of tfoR abolished competence for exogenous DNA even when (GlcNAc)2 was provided. The introduction of a plasmid clone carrying the tfoR+ gene into the tfoR deletion mutant complemented the competence deficiency. We also found that the tfoR gene encodes a 102-nucleotide small RNA (sRNA), which was transcriptionally activated in the presence of (GlcNAc)2. Finally, we showed that this sRNA activated translation from tfoXVC mRNA in a highly purified in vitro translation system. Taking these results together, we propose that in the presence of (GlcNAc)2, TfoR sRNA is expressed to activate the translation of tfoXVC, which leads to the induction of natural competence.


Natural competence is the ability of bacteria to actively take up DNA from the environment. The imported DNA is homologously integrated into the recipient chromosome to cause a gene exchange, called transformation, while being degraded into nucleotides to be used as a nutrient (15, 34). In most well-characterized naturally competent bacteria (except for constitutively competent bacteria, such as Neisseria gonorrhoeae and Neisseria meningitides [59]), the development of a competent state is tightly controlled (10), indicating that the advantages of DNA uptake depend on environmental changes.

Cameron et al. proposed that bacteria of the families Pasteurellaceae, Enterobacteriaceae, and Vibrionaceae might share a mechanism to regulate competence genes, whose expression is controlled by two activator proteins, TfoX (also known as Sxy) and CRP (5). Although competence genes are present in these families, only a few species are known to be naturally competent. Haemophilus influenzae (Pasteurellaceae), one of the well-characterized species, becomes moderately competent as growth becomes slow during late log phase in rich medium and becomes maximally competent when log-phase cells are transferred to a defined starvation medium (23). Under competence-inducing conditions, depletion of sugar imported by a phosphotransferase system causes a rise in cAMP levels, resulting in the formation of its active complex with CRP (cAMP-CRP), which, together with H. influenzae TfoX (TfoXHI), activates the transcription of competence genes (25, 35, 52). tfoXHI expression is activated at the transcriptional and translational levels in cells in a competence-inducing environment (6, 52). The involvement of the cAMP-CRP complex in transcriptional control has already been suggested (6, 77). Although the tfoXHI mRNA secondary structure plays a crucial role in translational control (6), the regulatory mechanism is not fully understood.

Vibrio cholerae, the agent of cholera, causes a dehydrating diarrheal illness. However, outside the human host, V. cholerae lives in natural aquatic environments, such as rivers, estuaries, and coastal waters, where it often associates with chitinaceous zooplanktons, such as copepods (32). Chitin, a polymer of β-1,4-linked N-acetylglucosamine (GlcNAc), not only serves as a surface for colonization in aquatic habitats (9, 42, 53) and as a carbon and nitrogen source (24, 30, 42, 49) but also induces natural competence (41). Blokesch and Schoolnik demonstrated that the whole O1-specific antigen gene cluster of V. cholerae O1 El Tor can be exchanged by the O37- or O139-antigen gene cluster through chitin-induced natural competence (4). In addition, the cholera toxin prophage (65) and clusters of metabolic genes (43) can be transferred in the same manner. Recently, other Vibrio species, Vibrio vulnificus, Vibrio fischeri, and Vibrio parahaemolyticus, were shown to be naturally transformable upon exposure to chitin (8, 17, 50). Thus, chitin-induced natural competence may be a common feature of different Vibrio spp. This mode of gene transfer seems to be limited to niches where chitin is abundant in the environment, for example, on the surface of copepods. Vibrio species are the most abundant bacterial species associated with copepods (55, 57), where there may be a pool of DNA from closely related species. Under such environment, chitin might serve as a signal for these bacteria to sense a DNA source suitable for genetic exchange via transformation.

In V. cholerae, TfoXVC (an ortholog of TfoXHI) plays a central role in chitin-induced competence (41, 42). The expression of the tfoXVC gene is activated in response to chitin, and TfoXVC upregulates competence genes, including VC1917, encoding an essential competence protein homologous to Bacillus subtilis ComEA, and structural genes for type IV pilus-like DNA uptake machinery (pilA, pilB, pilQ, etc). TfoXVC also positively regulates several genes (VCA0027, VC1952, VC0679, and VC0972) for extracytoplasmic chitin degradation functions (41). Our previous studies demonstrated that chitin, more specifically the disaccharide (GlcNAc)2, activates both the transcription and translation of tfoXVC to induce competence (71). Therefore, it is suggested that H. influenzae and V. cholerae use a similar mode of genetic control for competence development. In this study, we screened a multicopy library of V. cholerae DNA fragments and identified a novel small RNA (sRNA) involved in translational regulation of tfoXVC. This is the first report to identify a posttranscriptional regulator of the tfoX gene among naturally competent species.


Bacterial strains, plasmids, media, and DNA manipulation.

All the strains and plasmids used in this study are listed in Table 1. The Escherichia coli strain JM109 (72) was used for DNA cloning. The bacteria were grown in Luria-Bertani (LB) broth and agar and SOC (2% tryptone, 0.5% yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4, 20 mM glucose). The antibiotics used and their concentrations were as follows: 100 μg/ml of ampicillin (Ap), 12.5 μg/ml (for E. coli) or 1 μg/ml (for V. cholerae) chloramphenicol (Cm), and 30 μg/ml kanamycin. Arabinose, 5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside (X-Gal), and isopropyl-β-d-thiogalactopyranoside (IPTG) were used at concentrations of 5 mg/ml, 100 μg/ml, and 1 mM, respectively. PCR amplification of DNA was carried out with a GeneAmpPCR system 9700 (Applied Biosystems) by using the Herculase II fusion enzyme (Stratagene). DNA sequencing was performed using a BigDye terminator v1.1 cycle sequencing kit (Applied Biosystems). Customized primers, the nucleotide sequences of which are listed in Table S1 in the supplemental material, were purchased from Hokkaido System Sciences. Chromosomal and plasmid DNAs were extracted using a DNeasy blood and tissue kit (Qiagen) and a High Pure plasmid isolation kit (Roche), respectively.

Table 1.
Strains and plasmids used in this study

Protein analysis.

SDS-PAGE and Western blotting of proteins were performed according to previously described methods (70).

Chromosomal engineering.

Genetically modified E. coli and V. cholerae strains were constructed using the λ Red-FLP recombination system (12, 70, 71). Briefly, the procedure includes 3 processes: (i) preparation of a PCR fragment typically containing 2 Flp recognition target (FRT)-linked antibiotic resistance gene cassettes with flanking regions homologous to the target gene (X::FRT::cat::FRT::X′ or X::FRT::kan::FRT::X′ fragment, where X is the upstream homologous sequence and X′ is the downstream homologous sequence; the typical size of X or X′ is ~50 bp for E. coli or ~1,000 bp for V. cholerae); (ii) introduction of the PCR fragment into the recipient strain expressing λ Red recombinase to integrate the exogenous DNA into the chromosome; and (iii) elimination of the antibiotic resistance gene by FLP recombinase if necessary.

(i) Preparation of the PCR fragment to construct the blap::tfoXVC::lacZ translational fusion in E. coli.

The PCR fragment was generated by a 2-step PCR. pKD3 (12) was used as a template to amplify the FRT::cat::FRT cassette flanked by the 50-bp sequence homologous to the upstream region of the E. coli lac promoter and the 20-bp sequence homologous to the upstream region of the bla promoter (blaP) using the primers EClacI-FRTf1/FRT-blar1. On the other hand, the blaP fragment flanked by the 20-bp sequence homologous to tfoXVC (nucleotides +1 to +20) was amplified using the primers FRT-blaf1/bla-tfoXr1 and pKD3. Additionally, we amplified the tfoXVC fragment (nucleotides +1 to +709) flanked by the 50-bp sequence homologous to lacZ by using the primers bla-tfoXf1/tfoX-lacZr6 and chromosomal DNA from V060002. These 3 PCR fragments were purified using a High Pure PCR product purification kit (Roche). Then, 0.3 pmol of each of the three PCR fragments was mixed with the primers EClacI-FRTf1/tfoX-lacZr6. The PCR program was as follows: 95°C for 2 min, followed by 25 cycles at 95°C for 30 s, 40°C for 30 s, and 72°C for 2 min, which yielded the DNA fragment lacZ::FRT::cat::FRT::blaP::tfoXVC-TL::lacZ′.

(ii) Preparation of the PCR fragment for hfq gene disruption in E. coli.

The PCR fragment for hfq disruption was prepared as described previously (44). The 500-bp upstream and downstream regions flanking hfq were amplified using the primer sets hfqU1/hfqU2 and hfqD1/hfqD2. Additionally, the kan cassette was amplified from pTH18ks5 (21) using the primers Km1/Km2. These 3 PCR fragments were purified and then connected by PCR using the primers hfqU1/hfqD2 to obtain the DNA fragment hfq::kan::hfq′.

(iii) Preparation of the PCR fragment for tfoR gene disruption in V. cholerae.

The 2-step PCR protocol described in section i was used to generate the DNA fragment. Chromosomal DNA from V060002 was used as a template to amplify the 1,000-bp upstream and downstream regions flanking tfoR with the primer sets TFORf15/TFOR-FRTr1 and TFOR-FRTf1/TFORr1, respectively. Additionally, pKD4 (12) was used as a template to amplify the FRT::kan::FRT cassette with the primers FRTf1/FRTr1. These 3 PCR fragments were purified and then connected by PCR using the primers TFORf15/TFORr1 to obtain the DNA fragment tfoR::FRT::kan::FRT::tfoR′.

(iv) Preparation of the PCR fragment to construct the tfoR+78::lacZ transcriptional fusion in V. cholerae.

The PCR fragment for construction of the tfoR+78::lacZ transcriptional fusion was generated by a 2-step PCR. Chromosomal DNA from V060002 was used as a template to amplify the 1,000-bp upstream and downstream regions flanking tfoR with the primer sets TFORf15/TFORr10 and TFORf14/TFORr1, respectively. Additionally, we amplified the lacZ::FRT::cat::FRT fragment with the primers TFOR-LACZf3/TFOR-FRTr3 by using chromosomal DNA from SY0605, which contains a lacZ::FRT::cat::FRT cassette within the ctxA gene. These 3 PCR fragments were purified and then connected by PCR using the primers TFORf15/TFORr1. The PCR program was as follows: 95°C for 2 min, followed by 25 cycles at 95°C for 30 s, 40°C for 30 s, and 72°C for 4 min, which generated the DNA fragment tfoR+78::lacZ::FRT::cat::FRT::tfoR′.

(v) Preparation of the PCR fragment to construct the tfoR−30::lacZ transcriptional fusion in V. cholerae.

The 2-step PCR protocol described in section iv was used to amplify the DNA fragment. Chromosomal DNA from V060002 was used as a template to amplify the 1,000-bp upstream and downstream regions flanking tfoR with the primer sets TFORf15/TFORr13 and TFORf14/TFORr1, respectively. We also amplified the lacZ::FRT::cat::FRT fragment with the primers TFOR-LACZf5/TFOR-FRTr3 by using chromosomal DNA from SY0605. These 3 PCR fragments were purified and then connected by PCR using the primers TFORf15/TFORr1 to obtain the DNA fragment tfoR−30::lacZ::FRT::cat::FRT::tfoR′.

(vi) Recombination by λ Red and FLP.

In vivo recombination mediated by λ Red and FLP was conducted as described previously (70, 71). The PCR products were purified as mentioned above, ethanol precipitated, and dissolved in 20 μl of 10% glycerol. Each DNA (2 μg) was electroporated into the recipient E. coli and V. cholerae strains expressing λ Red recombinase. Table S2 in the supplemental material shows the specific combinations of donor DNA and recipient strain for the chromosomal engineering.

Construction of pRL-SYX1.

The fragment including the upstream sequence of tfoXVC (nucleotides −238 to +308) was PCR amplified from chromosomal DNA of V060002 using the primers TFOXPf1S/TFOXPr1B and then cloned into the SalI/BamHI site of pRL-124 (39) to generate pRL-SYX1.

Construction of pRL-SYX2, pRL-SYX3, pRL-SYX4, pRL-SYX5, pRL-SYX6, and pRL-SYX7.

The fragment including tfoXVC (nucleotides +1 to +308) fused to the tac promoter (tacp::tfoXVC) was PCR amplified from chromosomal DNA of SY0617 (71) using the primers PTRCf1S/TFOXPr1B and then cloned into the SalI/BamHI site of pRL-124 to generate pRL-SYX2. Using the same primers, a fragment of tacp::tfoXVC (nucleotides +93 to +308) was amplified from chromosomal DNA from SY0620 (71) and then inserted into the SalI/BamHI site of pRL-124 to yield pRL-SYX3. The same regions with a single point mutation were amplified by 2-step PCR. Using several primer sets, PTRCf1S/TFOXmtr1 and TFOXmtf1/TFOXPr1B for pRL-SYX4, PTRCf1S/TFOXmtr2 and TFOXmtf2/TFOXPr1B for pRL-SYX5, PTRCf1S/TFOXmtr4 and TFOXmtf4/TFOXPr1B for pRL-SYX6, and PTRCf1S/TFOXmtr18 and TFOXmtf18/TFOXPr1B for pRL-SYX7, the 4 fragment sets were first amplified from pRL-SYX3. These fragments were purified and then fused by PCR using the primers PTRCf1S/TFOXPr1B to be cloned into the SalI/BamRI site of pRL124, generating the respective plasmids (pRL-SYX2, pRL-SYX3, pRL-SYX4, pRL-SYX5, pRL-SYX6, and pRL-SYX7).

Constructions of pRL-SYX8 and pRL-SYX9.

The tacp::tfoXVC fragment (nucleotides +93 to +308) was PCR amplified from pRL-SYX3 using the primers PTRCf1S/TFOXPr1E and then cloned into the SalI/EcoRI site of pRL-124 to generate pRL-SYX8. The same region with a single point mutation was amplified using the same primers and the template pRL-SYX6 and then inserted into the SalI/EcoRI site of pRL-124, yielding pRL-SYX9.

Construction of pBAD-SYR1.

The tfoR gene was PCR amplified using the primers tfoRf6Sc/tfoRr3H and chromosomal DNA from V060002. The amplified product was digested with SacI and HindIII and inserted into pBAD33 (18) to generate pBAD-SYR1.

Construction of pTrc-SYQ1.

The hfq gene from V. cholerae was PCR amplified using the primers HFQf2E/HFQr2S and chromosomal DNA from V060002. The amplified product was digested with EcoRI and SalI and inserted into pTrc99A (3) to generate pTrc-SYQ1.

Library screening.

Chromosomal DNA from V060002 was sonicated and then blunt ended using an End-It DNA end repair kit (Epicentre) according to the manufacturer's instructions. The DNA fragments were cloned into the HincII site of pUC118 (72). Transformations into SYEC001S were performed by electroporation. Cells were plated on LB agar containing Ap and X-gal to produce about 300 colonies per plate and incubated at 37°C. About 20,000 colonies were screened, and a single blue colony was isolated. Plasmid DNA was extracted, and the inserted fragment was sequenced using the primer M13RV.

β-Galactosidase enzyme assay.

β-Galactosidase activity was assayed as described previously (71) using E. coli and V. cholerae cells grown under the following culture conditions with each sample assayed in triplicate.

(i) E. coli.

Cells were grown to an optical density at 600 nm of ~0.4 in LB at 37°C. If necessary, Cm, Ap, arabinose, or IPTG was added to the medium.

(ii) V. cholerae.

Cells were grown in M9 minimal medium containing minimal essential medium vitamin solution (Gibco) and 5 mM GlcNAc or (GlcNAc)2 (Seikagaku Biobusiness) as described previously (71). If necessary, Ap was added to the medium.

Natural transformation.

Natural transformation was performed using the recipient V. cholerae cells grown in M9 minimal medium containing minimal essential medium and 5 mM GlcNAc or (GlcNAc)2 and chromosomal DNA from SY1001, which harbors a cat cassette in the ctxB gene, as described previously (71). If necessary, Ap was added to the medium. Transformation efficiency was defined as Cm-resistant (or Cm-and Ap-resistant) CFU divided by total viable (or Ap-resistant) CFU. Experiments were done at least twice.

Synthesis of digoxigenin-labeled RNA probe for Northern blotting.

The RNA probe complementary to tfoR was prepared as follows. The DNA fragment with the T7 promoter sequence was PCR amplified using the primers T7-tfoRr1/tfoRf3 and chromosomal DNA from V060002. This amplified DNA was used as a template for in vitro RNA synthesis from T7 RNA polymerase with a digoxigenin (DIG) RNA labeling kit (Roche). The DIG-labeled RNA was precipitated by ethanol and dissolved in diethylpyrocarbonate-treated H2O.

Northern blotting.

Cells were cultured under the same conditions used in β-galactosidase assays. RNAs were extracted from the cells using Isogen (Wako) according to the manufacturer's instructions. The RNAs (1.5 μg) were heated and cooled quickly; the RNAs were then separated electrophoretically on a 6% polyacrylamide gel containing 6 M urea. RNA Century-plus size markers (Ambion) were also separated as a molecular standard on the same gel. After electrophoresis, RNAs were electrotransferred onto a Hybond-N+ nylon membrane (GE Healthcare). After UV cross-linking, the membrane was treated for 2 h at 65°C in hybridization buffer (DIG Easy Hyb; Roche). Then, the membrane was incubated for 12 h at 65°C in hybridization buffer containing DIG-labeled RNA probe. After hybridization, the membrane was first washed under a low-stringency condition in 2× SSC (1×SSC is 0.15 M NaCl plus 0.015 M sodium citrate [pH 7.0]) containing 0.1% SDS for 10 min at room temperature and then under a high-stringency condition in 0.2× SSC containing 0.1% SDS for 30 min at 65°C. The membrane was treated with maleic acid buffer (0.1 M maleic acid [pH 7.5], 0.15 M NaCl) containing 1% blocking reagent (Roche), incubated for 20 min with 10,000-fold-diluted anti-DIG-AP Fab fragments (Roche), and then washed twice with maleic acid buffer containing 0.3% Tween 20. DIG-labeled signals were amplified by hydrolysis of disodium 3-(4-methoxyspiro{1,2-dioxetane-3.2′-(5′-chloro)tricyclo[,7]decan}-4yl)phenylphosphate (Roche) in detection buffer (0.1 M Tris-HCl [pH 9.5], 0.1 M NaCl) and visualized on a Hyperfilm ECL film (GE Healthcare).

Circularization RT-PCR.

Circularization reverse transcription (RT)-PCR (11, 73) was performed to determine the 5′ and 3′ ends of TfoR sRNA as follows. The total RNA used in Northern blotting [isolated from cells cultured in the presence of (GlcNAc)2] was also used for self-ligation. Prokaryotic mRNA has a triphosphate at its 5′ terminus. To render the 5′ terminus ligatable, the triphosphate was removed by phosphatase and then monophosphate was added to the 5′-OH end by using a KinaseMax kit (Ambion). Then, the processed RNA (8 μg) was subjected to a reaction with T4 RNA ligase (Takara). cDNA was reverse transcribed from 4 μg of ligated RNA with the primer tfoR-CRTr1, using avian myeloblastosis virus (AMV) reverse transcriptase XL (Takara) according to the manufacturer's instructions. The 5′-3′ junction region in the cDNA was PCR amplified using the primers TFORr5/tfoR-CRTf1. The major products obtained (Fig. 3B) were gel purified and cloned into pGEM-T (Promega). Seven clones from independent transformants were sequenced using the primer T7P.

Fig. 3.
Identification of the tfoR gene. (A) Intergenic region between the VC2078 and VC2079 genes. The cloned fragment in pUC-SYR1 is indicated by the shaded arrow that points in same direction as the lac promoter. The tfoR gene revealed by circularization RT-PCR ...

Synthesis of RNAs for in vitro translation.

DNA templates carrying a T7 promoter sequence were generated by PCR using chromosomal DNA from V060002 and specific primers. For TfoR (nucleotides +1 to +102) and its cRNA, the primer sets T7-tfoRf6/tfoRr3 and T7-tfoRr1/tfoRf17, respectively, were used. For transcription templates generating tfoXVC::flag RNAs, full-length RNA (nucleotides +1 to +709 translationally fused to the FLAG tag sequence), and ΔTLE RNA (nucleotides +93 to +709 translationally fused to the FLAG tag sequence), we performed a 2-step PCR: the first round of PCR was carried out using the primers T7-TFOXUTRf42/TFOX-2FLAGr1 for full-length RNA and the primers T7-TFOXUTRf6/TFOX-2FLAGr1 for ΔTLE RNA; the second round of PCR was done using the primers T7-TFOXUTRf42/2FLAG-T7Tr1 for full-length RNA and the primers T7-TFOXUTRf6/2FLAG-T7Tr1 for ΔTLE RNA. Using each amplified DNA as the template, RNA was synthesized by T7 RNA polymerase with a MAXIscript T7 kit (Ambion). After the reaction, the template DNA was degraded by Turbo DNase (Ambion) and the synthesized RNA was purified using a MEGAclear kit (Ambion).

In vitro translation.

An in vitro translation reaction was performed in a highly purified system (PURE system, purchased from New England BioLabs as Purexpress) (54). The translation mixture (25 μl) contained 40 nM tfoXVC::flag RNA and various concentrations (0, 40, 200, and 400 nM) of TfoR. After incubation for 20 min at 37°C, the synthesized TfoXVC::FLAG proteins were detected by Western blotting using a monoclonal anti-FLAG M2-horseradish peroxidase antibody (Sigma).

RNAhybrid analysis.

The RNAhybrid software program (26) was run on the website using the sequences of tfoR (nucleotides +1 to +102) and tfoXVC (nucleotides +43 to +133).

BLAST analysis.

The sequences homologous to tfoR were searched using BLAST (1) on the web site The genome sequences used were from V. cholerae N16961 (accession no. AE003852.1), V. parahaemolyticus 2210633 (accession no. BA000031.2), V. vulnificus CMCP6 (accession no. AE016795.3), and V. fischeri ES114 (accession no. CP000020.2).

Multiple alignment.

The alignment among the tfoR homologues was done using CLUSTAL W (62) in the GENETYX-MAC version 12.0.9 software program (Genetyx Corporation).

Nucleotide sequence accession number.

The tfoR sequence was deposited in GenBank and assigned the accession number AB598400.


Reevaluation of DSE in tfoXVC.

Previously, we identified 2 cis-acting elements for translational control of tfoXVC, a translational control element (TLE) nucleotides +43 to +84) and a downstream element (DSE) (nucleotides +107 to +133) (71) (Fig. 1A). TLE negatively controls translation in the absence of (GlcNAc)2, whereas DSE seems to play an essential role in translation. DSE is located downstream of a putative Shine-Dalgarno (SD) sequence (GGGA, +98 to +101) and start codon (GTG, +104 to +106), which are predicted on the basis of the GenBank sequence annotation (accession number AE003852.1), and thereby it has been postulated to be a translational enhancer, such as a downstream box (60, 71). By careful reannotation, however, we found that there is another conservative SD sequence (AGGA, +112 to +115) located with other start codons (ATG, +119 to +121; ATG, +125 to +127) within DSE (Fig. 1A). This suggests a possibility that DSE overlaps the translational start region of tfoXVC. To clarify this, we introduced a point mutation in each potential start codon within a tfoXVC::lacZ translational fusion on the plasmid pRL124. We first constructed a fusion plasmid, pRL-SYX3, in which both the native promoter (tfoXVCp) and TLE were exchanged with the tac promoter (tacp) to constitutively drive transcription and translation (Fig. 2). This enabled us to exclude a possibility that point mutation may affect the regulation of tfoXVC expression. Three other plasmids (pRL-SYX4, pRL-SYX5, and pRL-SYX6) (Fig. 2) were constructed from pRL-SYX3. Derivatives of V. cholerae strain V060002 containing a fusion plasmid with or without a single base change were tested for β-galactosidase activity when they were grown in M9 minimal medium in the presence of GlcNAc or (GlcNAc)2 (Fig. 2). Consistent with our previous data using chromosomal fusion (71), the addition of (GlcNAc)2 increased β-galactosidase activity from strains carrying pRL-SYX1 (15-fold) and pRL-SYX2 (21-fold), which contain the wild-type leader sequence situated downstream of tfoXVCp and tacp, respectively, compared with that of GlcNAc, while the strain carrying pRL-SYX3 had a high activity of β-galactosidase under both culture conditions (Fig. 2). A similar pattern of expression was observed in strains carrying pRL-SYX4 and pRL-SYX5, which have base changes of GTG at positions +104 to +106 (GTG+104 to +106) to CTG and ATG+119 to +121 to CTG, respectively (the underlined nucleotide was changed), indicating that these nucleotides are not essential for translation. On the other hand, β-galactosidase activity in the strain with pRL-SYX6, in which ATG+125 to +127 was replaced by CTG, was reduced nearly to a background level under both culture conditions (Fig. 2). We also changed the A+125 to C in the tacP/ΔTLE-based transcriptional fusion (pRL-SYX8) to generate pRL-SYX9 (Fig. 2). The two strains carrying pRL-SYX8 and pRL-SYX9 showed similar levels of expression (Fig. 2). These results indicated that the A+125-to-C mutation impaired translation but not transcription. Thus, it appears plausible that ATG+125 to +127 acts as the start codon, rather than GTG+104 to +106 (Fig. 1A). According to the position of the start codon, the SD sequence could be comprised of AGGA+112 to +115 (Fig. 1A). Consistent with this, the base change of AGGA+112 to +115 to ACCA (pRL-SYX7) decreased β-galactosidase activity (Fig. 2). This notion is further supported by a previous report in which optimal spacing between the SD sequence and the start codon is described as about 8 to 10 nucleotides (nt) (7).

Fig. 1.
(A) Mfold prediction of secondary structure of tfoXVC mRNA (71). Positions corresponding to TLE and DSE are drawn in gray. Possible base pairs are indicated by bars. Putative start codon and SD sequence for tfoXVC are shown by bold letters. Of these, ...
Fig. 2.
Determination of the translational initiation region of tfoXVC. Left: the structures of the tfoXVC::lacZ fusions on pRL124. Names of plasmids are indicated on the left. The upstream and coding regions of tfoXVC are shown in gray. tfoXVCp and tacp indicate ...

Genetic screening for the positive regulator of tfoXVC.

We previously predicted an intramolecular base pairing between the TLE and DSE RNAs using the Mfold software program (71, 76). When this structure was reevaluated using the information as described above (Fig. 2), TLE seemed to sequester the SD sequence within DSE (Fig. 1A). Based on these observations, we proposed a model for the translational control mechanism as follows. In the absence of (GlcNAc)2, the TLE of the tfoXVC mRNA represses translation through base pairing with the DSE of the mRNA. On the other hand, in the presence of (GlcNAc)2, the formation of this structure is somehow inhibited by an unknown substance, allowing the ribosome to bind to the site of DSE, which ultimately enhances translation. If this model is true, some regulatory factor(s) may be needed to activate translation of tfoXVC in response to (GlcNAc)2. Therefore, we thought that it might be possible to clone the regulatory gene(s) by screening for increased translational activity of tfoXVC. To achieve this, we constructed an E. coli strain, DH1 (20), with a tfoXVC::lacZ translational fusion (SYEC001S) whose transcription was constitutively driven by the bla promoter from Tn3 (14). SYEC001S was transformed with a plasmid library consisting of sonicated DNA fragments from a V. cholerae strain, V060002, cloned into the HincII site of pUC118. The resultant colonies were screened on LB plates containing X-Gal and Ap. On these plates, the starting strain appeared white (Lac). Among 20,000 colonies screened, a single blue colony (Lac+) was isolated, which carried the plasmid pUC-SYR1. Retransformation of SYEC001S with pUC-SYR1 confirmed that this phenotype was a plasmid-linked trait. pUC-SYR1 increased β-galactosidase activity from the tfoXVC::lacZ fusion in SYEC001S more than 30-fold (145 ± 18 Miller units [mean ± standard deviation] in LB) compared with that of pUC118 (4 ± 3 Miller units). These results suggest that pUC-SYR1 carries a positively trans-acting regulatory element of tfoXVC. Sequence analysis showed that pUC-SYR1 contained the 332-bp intergenic region between the VC2078 and VC2079 genes (corresponding to nucleotide positions 2236523 to 2236854 on chromosome I in the V. cholerae strain N16961) (Fig. 3A). Computer analysis of the cloned fragment using GENETIX predicted no significant open reading frames. Instead, this fragment contained an apparent Rho-independent terminator oriented in the 3′ direction of the lac promoter and was already predicted to encode a 96-nt RNA (corresponding to nucleotide positions 2236607 to 2236702 on chromosome I in N16961) by an RNA deep-sequencing-based screen (33). These observations suggest that pUC-SYR1 might express a noncoding regulatory RNA. We called this gene in the 332-bp region “tfoR.”

Effect of tfoR on tfoXVC expression in V. cholerae.

To examine the effect of tfoR on tfoXVC expression in V. cholerae, we introduced a deletion mutation of tfoR into the strain V060002 with a tfoXVC::lacZ transcriptional or translational fusion and assayed β-galactosidase activities from these strains grown in the presence of GlcNAc or (GlcNAc)2 (Table 2). As described previously (71), in the tfoR+ background, the addition of (GlcNAc)2 moderately (2-fold) activated the transcriptional fusion gene but strongly (21-fold) activated the translational fusion gene compared with that of GlcNAc (Table 2). However, in the ΔtfoR background, the expression of the translational fusion was scarcely increased by (GlcNAc)2 (Table 2). Deletion of tfoR had no significant effect on the expression of the transcriptional fusion (Table 2). In addition, pUC-SYR1 stimulated the expression of the translational fusion (10- to 14-fold) independently of (GlcNAc)2 compared with pUC118 (Table 2, compare tfoXVC-TL::lacZΔtfoR/pUC118 and tfoXVC-TL::lacZΔtfoR/pUC-SYR1). These results suggest that tfoR positively affects tfoXVC expression, mainly at the translational level, and multicopy tfoR bypasses the need of (GlcNAc)2 for translational regulation.

Table 2.
Effect of tfoR on expression of tfoXVC::lacZ fusions in V. cholerae

Effect of tfoR on natural competence.

The data presented above revealed tfoR-dependent translational activation of tfoXVC, encoding a positive regulator of natural competence. Therefore, we tested the effect of tfoR on transformation. After the competence-inducing treatment, the wild-type strain was transformed with a frequency of 1.3 × 10−8 (Table 3). However, the tfoR mutant strain was not transformed under the same culture conditions (Table 3). The introduction of pUC-SYR1 carrying the tfoR+ gene into the tfoR mutant induced transformation regardless of the presence or absence of (GlcNAc)2, confirming that tfoR was required for competence. Furthermore, pUC-SYR1 did not stimulate transformation in the tfoXVC mutant (Table 3). This indicates that the positive effect of tfoR on competence is mediated through tfoXVC. It was noted that the tfoR mutant strain with pUC-SYR1 had a much higher efficiency of transformation (68- to 330-fold) than the wild-type strain (Table 3), although translation activity of tfoXVC in the strain with pUC-SYR1 was slightly lower (2- to 2.5-fold; for example, compare the activities of 317 ± 10 units and 129 ± 5 units in Table 2) than the activity from the wild-type strain grown in the presence of (GlcNAc)2 (Table 2). This discrepancy is discussed in the following section.

Table 3.
Effect of tfoR on natural transformation

Identification of a novel RNA activator.

To detect RNA product from tfoR, we performed Northern blotting by using a tfoR-specific RNA probe. This probe was created to detect the transcript that ended at the site of a putative Rho-independent terminator. Total RNA was isolated from cells grown under the same culture conditions for β-galactosidase assays. In the presence of (GlcNAc)2, an ~100-nt RNA was detected from the total RNA prepared from the wild-type cells, whereas no bands were observed in the total RNA from the tfoR mutant cells (Fig. 4). We could also detect this sRNA in the tfoXVC mutant (Fig. 4). Furthermore, when the RNA probe for the opposite transcript was used, the sRNA was not detected (data not shown). The tfoR mutant strain with pUC-SYR1 produced independently of (GlcNAc)2 much larger amounts of the 100-nt RNA than the wild-type strain (Fig. 4), which correlated with a remarkable increase in transformation efficiency (Table 3). However, the introduction of pUC-SYR1 into the tfoR mutant was not much more effective for tfoXVC expression than the addition of (GlcNAc)2 to the wild-type strain (Table 2). The difference between a multicopy effect of tfoR on tfoXVC and on competence may suggest an additional positive role of tfoR in competence beyond the tfoXVC regulation. Additionally, we detected several longer RNAs (200 to 500 nt), as well as the 100-nt RNA in the strain carrying pUC-SYR1 (Fig. 4). As expected from their sizes, the longer species may be hybrid transcripts that contain both sequences of tfoR and pUC118.

Fig. 4.
(GlcNAc)2-dependent expression of TfoR sRNA. Total RNA was isolated from cells grown in the presence of GlcNAc or (GlcNAc)2, separated on a polyacrylamide gel containing 6 M urea, and transferred onto a nylon membrane. RNA product from tfoR was detected ...

The 5′ and 3′ ends of the (GlcNAc)2-induced sRNA were determined by circularization RT-PCR (11, 73). Briefly, the 3′ and 5′ ends of a linear RNA molecule were joined by T4 RNA ligase, and a region containing the 3′-5′ junction was converted to cDNA by reverse transcriptase. The cDNA was successively amplified by PCR, with one primer close to the 3′ end and the other to the 5′ end. Cloning and sequencing of amplified DNA fragments would definitively demonstrate the number and species of nucleotides between the 3′ and 5′ ends of a given transcript. After ligation, total RNA was subjected to reverse transcription and synthesized cDNA was utilized as a template for PCR. The major band (slightly less than 100 bp) of PCR products (Fig. 3B) was cloned into pGEM-T and transformed in E. coli, and 7 clones from independent transformants were sequenced. All clones exhibited the same 5′ end, “A” (Fig. 3A). This established that the 5′ end of the sRNA was the A nucleotide indicated as +1, which was located 1 nt downstream of the 5′ end of the sRNA previously predicted by Liu et al. (33). From the sequence result of 6 among 7 analyzed clones, it was indicated that the 3′ end of sRNA was the last “U” of 7 U's following the terminator stem (Fig. 3A), which was suggested to be the major transcription termination site. Based on these results, we estimated the sRNA to be 102 nt long. This is consistent with the size of the sRNA obtained from Northern blotting (Fig. 4).

We could not find a significant promoter immediately upstream of the 5′ end of the sRNA. To evaluate the transcriptional response of the sRNA gene to (GlcNAc)2, we fused the lacZ gene to a site within the terminator sequence (immediately downstream of nucleotide +78) (Fig. 3A). The expression of the fusion gene was induced 16-fold by (GlcNAc)2 (Table 4). Moreover, we inserted lacZ into a further upstream nucleotide (immediately downstream of nucleotide −30) (Fig. 3A). The fusion gene expression still responded to (GlcNAc)2 (Table 4), suggesting that the (GlcNAc)2-responsive promoter may be located further upstream of the −30 nucleotide G.

Table 4.
Analysis of tfoR::lacZ fusion expression in V. cholerae

We cloned this sRNA gene (nucleotides +1 to +102) in pBAD33. The expression of the RNA was inducible under the control of the araBAD promoter. This plasmid, pBAD-SYR1, was used to transform SYEC001S, an E. coli strain with a tfoXVC::lacZ translational fusion gene. Induction of transcription of the sRNA gene in this plasmid was necessary and sufficient to stimulate the expression of the tfoXVC::lacZ translational fusion (Table 5). We concluded that this sRNA gene is tfoR, and we called this novel sRNA “TfoR.”

Table 5.
Effect of tfoR and hfq on expression of the tfoXVC::lacZ translational fusion in E. coli

Effect of Hfq on TfoR-mediated regulation of tfoXVC.

The majority of bacterial sRNA regulators act through base pairing with RNAs, resulting in the modulation of translation and stability of mRNAs (68). The RNAhybrid program (26) was used to search for a possible complementary sequence in the 5′ untranslated region (UTR) of tfoXVC mRNA with the sequence of TfoR. TfoR was found to extensively match with the region including TLE of tfoXVC mRNA (Fig. 1B), which acts negatively on translation (71). Therefore, TfoR might contribute to the activation of translation through intermolecular base pairing with TLE. In many cases, the RNA chaperone Hfq is required for sRNA-mediated regulation to affect the processing or stability of the sRNA transcript (40, 46, 75) or to facilitate RNA-RNA interactions of sRNA and target mRNA (46, 74). To examine whether Hfq affects the translational regulation of tfoXVC by TfoR, we introduced a deletion mutation of hfq into the V060002 background. However, the hfq mutation severely impaired growth in M9 minimal medium containing (GlcNAc)2 (data not shown). For such a methodological limitation in V. cholerae, we reconstituted an experimental system based on the E. coli reporter strain SYEC001S to analyze the Hfq effect. We first deleted the hfq gene in SYEC001S to generate strain SYEC001SQ1, and then tfoXVC::lacZ expression was analyzed when TfoR and Hfq were expressed under the control of the araBAD promoter of pBAD33 and the tac promoter of pTrc99A, respectively. In the hfq mutant, either induction of TfoR from pBAD-SYR1 or Hfq from pTrc-hfq and pTrc-SYQ1 (carrying the E. coli hfq gene and the V. cholerae hfq gene, respectively) failed to stimulate translation, while both induction of TfoR and of Hfq restored the capability for activating translation (Table 5). Northern blotting showed that TfoR was detectable only from the hfq+ strains (Fig. 5), suggesting that Hfq may be required for TfoR expression.

Fig. 5.
Effect of hfq on TfoR expression in E. coli. Total RNA was isolated from cells carrying pBAD33 (or pBAD-SYR1) and pTrc99A (or pTrc-hfq/pTrc-SYQ1) grown in LB containing Cm, Ap, arabinose, and IPTG, separated on a polyacrylamide gel containing 6 M urea, ...

TfoR activates translation from tfoXVC mRNA in vitro.

For in vitro translation of tfoXVC, we used a reconstituted PURE system that includes only purified ribosomes and all other translation factors (54). This system is successfully used elsewhere for in vitro translation analysis (38, 48, 66). We in vitro synthesized tfoXVC::flag mRNA (full length: nucleotides +1 to +709 translationally fused to the FLAG tag sequence) as a template and TfoR sRNA. An RNA complementary to TfoR was also synthesized as a negative control. The TfoXVC::FLAG protein was analyzed by Western blotting with anti-FLAG antibody. TfoXVC::FLAG was synthesized in the presence of TfoR at an equimolar concentration of the template RNA, whereas the addition of the RNA complementary to TfoR at up to a 10-fold molar excess over the RNA template did not stimulate TfoXVC::FLAG synthesis (Fig. 6). This indicates that TfoR is required specifically for translational activation of tfoXVC. We also synthesized tfoXVC::flag mRNA with deletion of TLE (ΔTLE: nucleotides +93 to +709). The deletion made the translation constitutively active in vivo (71). Consistently, the TfoXVC::FLAG protein was synthesized from the ΔTLE mRNA even in the absence of TfoR, which was not further activated by the addition of TfoR (Fig. 6). This suggests that the TLE of the tfoXVC mRNA could be a regulatory target of TfoR.

Fig. 6.
Effect of TfoR on translation from tfoXVC mRNA in vitro. Template tfoXVC::flag mRNAs (full-length, nucleotides +1 to +709, and ΔTLE, nucleotides +93 to +709, both of which are translationally fused to the FLAG tag sequence) and TfoR sRNA were ...

TfoR is conserved in Vibrio spp.

In addition to V. cholerae, several Vibrio spp., including V. vulnificus, V. fischeri, and V. parahaemolyticus, show natural competence in the presence of chitin (8, 17, 50). These and many other Vibrio spp. have not only the tfoX gene but also DNA uptake genes (31, 50), suggesting a conservation of the TfoX regulon for competence induction. Using the BLAST program (1), we found homologues of V. cholerae tfoR (tfoRVC) in many sequenced genomes of the Vibrio spp. Of these, Fig. 7 shows the sequences homologous to that of tfoRVC from known competent species as above. In every case, the tfoR locus was situated between a gene encoding ferrous iron transport protein A (VC2078, VP0856, VV1_0149, and VF_0833) and a gene encoding a hypothetical protein homologous to an AraC/XylS-type transcriptional regulator (VC2080, VP0854, VV1_0150, and VF_0832) (Fig. 7A). These genomic features may indicate functional relatedness within the gene cluster. Multiple alignment analysis (62) showed a high similarity of tfoR (Fig. 7B). Furthermore, using the RNAhybrid program, we could predict significant base pairing of tfoR and tfoX in each species (data not shown). The presence of both tfoR and tfoX in various Vibrio spp. suggests the potential for a conserved mechanism of competence regulation across this genus.

Fig. 7.
Conservation of TfoR in the Vibrio spp. (A) The genomic location of the tfoR gene. Bacterial strains shown are as follows: Vc, V. cholerae N16961; Vp, V. parahaemolyticus 2210633; Vv, V. vulnificus CMCP6; Vf, V. fischeri ES114. In the genome sequences ...


TfoX is the central regulator of DNA uptake by the naturally competent bacteria H. influenzae and V. cholerae. Under competence-inducing conditions, tfoX gene expression is activated at the transcriptional and translational levels in both organisms (6, 71), but their precise regulatory mechanisms have remained unknown. We have investigated the mechanism of tfoXVC translational control in V. cholerae, which is more prominent than the transcriptional control. In this study, we identified a novel sRNA, TfoR sRNA, by a random screening of clones that are able to stimulate tfoXVC::lacZ translation (Fig. 3; see also “Genetic screening for the positive regulator of tfoXVC” above). We showed that TfoR activated tfoXVC translation both in vivo and in vitro (Tables 2 and and55 and Fig. 6) and was required for natural competence (Table 3). In addition, transcription of tfoR was induced by (GlcNAc)2 (Table 4 and Fig. 4). On the basis of these results, we propose that in the presence of (GlcNAc)2, TfoR sRNA is expressed to activate the translation of tfoXVC, which leads to the induction of natural competence. This is the first report of the identification of a posttranscriptional regulator for the expression of the tfoX gene. Regarding translational control in these H. influenzae and V. cholerae bacteria, however, the mechanisms are likely to be different, because there is no considerable sequence similarity between the 5′ UTRs of tfoXVC and tfoXHI (data not shown), and H. influenzae has no homologue of TfoR. On the other hand, Vibrio spp. seem to share the tfoR gene (Fig. 7) as well as the tfoX gene (50). This suggests that the tfoR/tfoX regulatory system may be important for controlling natural competence in Vibrio spp.

Many bacterial sRNA regulators act through base pairing with RNAs, resulting in the modulation of translation and stability of mRNAs (68). They can often base pair at the SD sequence and thus block translation initiation by preventing ribosome binding (68). In addition to repressing translation, sRNAs can also activate translation, in many cases by preventing the formation of an intrinsic structure inhibitory to translation initiation (19, 37, 47, 51, 66), which is called an “antiantisense mechanism” (16, 37). Using the RNAhybrid program, TfoR was predicted to extensively pair with the region including TLE of tfoXVC mRNA (Fig. 1), which acts negatively on translation (71). Therefore, TfoR might act as an antiantisense RNA through intermolecular base pairing with TLE to counteract its translational inhibitory effect. In many cases, sRNAs require the RNA chaperone Hfq for processing or stability (40, 46, 75) and for productive base pairing with target mRNAs (46, 74). In our V. cholerae strain background, disruption of hfq showed a phenotype with slow growth in M9 minimal medium containing (GlcNAc)2, suggesting that this mutation might affect utilization of (GlcNAc)2. Instead, our in vivo analysis in E. coli revealed the following: (i) that both Hfq and TfoR are required for activating translation of tfoXVC (Table 5), and (ii) that TfoR expression is remarkably decreased in the absence of Hfq (Fig. 5). On the other hand, TfoR could activate translation in vitro (PURE system) without adding the Hfq protein (Fig. 6). In order to explain the conflicting results, one possibility suggested is that Hfq affects translation of tfoXVC in vivo through modulating stability of TfoR. However, we cannot rule out the additional possibility that Hfq directly facilitates interactions of TfoR and tfoXVC RNAs. Therefore, further in vivo and in vitro analyses will be necessary to understand the role of Hfq in the TfoR regulation.

We have yet to determine the mechanism underlying TfoR expression. We found that the transcription of tfoR was significantly induced by (GlcNAc)2 (Table 4). Such a transcriptional response still occurred even though the lacZ reporter gene was fused immediately downstream of nucleotide −30 (Table 4). This suggests that a promoter that is directly or indirectly responsive to (GlcNAc)2 might be located further upstream of the position −30. In the multiple alignment analysis in Fig. 7B, a conserved sequence (TATAGT) similar to the −10 box of the σ70 promoter could be found at the nucleotides −42 to −37. If this acted as the promoter, longer RNA (at least 130 nt) would be primarily transcribed from it and then posttranscriptionally processed to yield the mature TfoR sRNA (102 nt). In this study, however, we could not detect such a longer transcript (Fig. 4). More extensive analysis will be required to elucidate the mechanisms of TfoR expression.

sRNA regulators have been identified in a wide range of bacteria and found to play critical regulatory roles in many physiological processes, for example, replication (63), outer membrane protein synthesis (45), sugar transport (67), stress response (2), iron homeostasis (40), virulence (47), and quorum sensing (29). In V. cholerae, several sRNAs have been identified through genetic screens and computational methods (13, 29, 58). Additionally, direct cloning and parallel sequencing experiments identified numerous sRNAs with unknown functions in this bacterium (33). In many competent bacteria, the development of a competent state is tightly controlled depending on environmental changes (10). This may mean that bacteria limit the incidence of genetic exchange caused by DNA uptake under unfavorable conditions. In both transformable species, Streptococcus pneumoniae and B. subtilis, competence is induced by a small peptide pheromone (22, 36). A dedicated two-component regulatory system detects the pheromone and transmits a signal, which leads to activation of the key transcriptional regulator for the competence regulon (10, 28, 64). On the other hand, H. influenzae utilizes the cAMP-CRP system to activate transcription of the tfoXHI gene (6, 77). To our knowledge, the present study is the first example of the involvement of sRNA in competence regulation. sRNAs are faster to produce than proteins and act posttranscriptionally. Thus, they could shut off or turn on gene expression more rapidly than protein-based transcriptional regulators. It is probable that sRNA-mediated control of tfoXVC expression confers an advantage to V. cholerae cells in quickly adjusting DNA uptake according to environmental changes. TfoXVC acts not only as a central regulator of the competence regulon but also as a positive regulator of several genes for extracytoplasmic chitin degradation (41), and thus it could promote degradation of chitin into oligosaccharides, including (GlcNAc)2, a major inducer of tfoXVC expression, generating a positive feedback loop. Therefore, it is reasonable to assume that TfoR may accelerate this regulatory loop via posttranscriptional activation of tfoXVC.

In addition to tfoXVC (VC1153), V. cholerae has another tfoX paralogue, tfoYVC (VC1722) (50), which was originally named tfoXGEMM (69). Both orthologs of tfoXVC and tfoYVC are present in other Vibrio spp., V. parahaemolyticus, V. vulnificus, and V. fischeri (50). Interestingly, the 5′ UTR of tfoYVC mRNA is reported to function as a cyclic di-GMP-sensing riboswitch to posttranscriptionally control its expression (27, 56, 61). Disruption of tfoYVC had no significant effect on chitin-induced transformation (S. Yamamoto, unpublished data); however, this gene might be involved in competence induction under some favorable environmental conditions. If this is the case, V. cholerae (and probably other Vibrio spp.) utilizes different posttranscriptional control mechanisms of paralogous regulators to regulate competence. Such regulatory modes of tfoXVC and tfoYVC might allow this bacterium to respond rapidly to different conditions required for DNA uptake.

Supplementary Material

[Supplemental material]


This work was supported in part by grants-in-aid from the Ministry of Health, Labor and Welfare of Japan (H22-Shokuhin-Ippan-012, H21-Shokuhin-Ippan-005, H21-Shokuhin-Ippan-013, H20-Shinko-Ippan-013, and H20-Shinko-Ippan-015) and the Ministry of the Environment of Japan (Global Environment Research Fund S-8).


Supplemental material for this article may be found at

[down-pointing small open triangle]Published ahead of print on 11 February 2011.


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