The Vibrio parahaemolyticus Small RNA RyhB Promotes Production of the Siderophore Vibrioferrin by Stabilizing the Polycistronic mRNA

Abstract

High-affinity iron acquisition in Vibrio parahaemolyticus is mediated by the cognate siderophore vibrioferrin. We have previously reported that the vibrioferrin biosynthesis operon (pvsOp) is regulated at the transcriptional level by the iron-responsive repressor Fur (T. Tanabe, T. Funahashi, H. Nakao, S. Miyoshi, S. Shinoda, and S. Yamamoto, J. Bacteriol. 185:6938–6949, 2003). In this study, we identified the Fur-regulated small RNA RyhB and the RNA chaperone Hfq protein as additional regulatory proteins of vibrioferrin biosynthesis. We found that vibrioferrin production was greatly impaired in both the ryhB and hfq deletion mutants, and a TargetRNA search (http://snowwhite.wellesley.edu/targetRNA/index2.html) revealed that the 5′-untranslated region of pvsOp mRNA (pvsOp 5′-UTR) contains a potential base-pairing region required for the formation of the RyhB-pvsOp 5′-UTR duplex. An electrophoresis mobility shift assay indicated that RyhB can directly bind to the pvsOp 5′-UTR with the aid of Hfq. Rifampin chase experiments indicated that the half-life of pvsOp mRNA in the ryhB and hfq mutants was approximately 3-fold shorter than that in the parental strain, suggesting that both RyhB and Hfq are engaged in the stabilization of pvsOp mRNA. Chrome azurol S assays followed by electrophoresis mobility shift assays and rifampin chase experiments carried out for mutant strains indicated that base pairing between RyhB and the pvsOp 5′-UTR results in an increase in the stability of pvsOp mRNA, thereby leading to the promotion of vibrioferrin production. It is unprecedented that RyhB confers increased stability on a polycistronic mRNA involved in siderophore biosynthesis as a direct target.

INTRODUCTION

Iron is an essential element for microbial survival and growth, because it serves as a component of heme and iron sulfur centers for crucial enzymes engaged in many key redox reactions. Although iron is the fourth most abundant element in the Earth's crust, its availability for bacteria is limited, because iron is usually present only in an extremely insoluble form, such as ferric hydroxide polymers under aerobic conditions at neutral pH. In mammalian hosts, iron is mostly sequestered in transferrin and lactoferrin as the first line of host defense against microbial pathogens (1). Hence, most bacteria possess specific systems to acquire iron under iron-depleted conditions. The most well-known strategy is a high-affinity iron transport system mediated by siderophores, which are low-molecular-weight ferric iron-binding chelators produced by microbes (2). In Gram-negative bacteria, ferric iron-loaded siderophore complexes (ferric siderophores) formed outside the cell are imported into the cell via a TonB-dependent specific uptake system comprising an outer membrane receptor and an ABC (ATP-binding cassette) transporter (3, 4). On the other hand, excessive intracellular free iron participates in Fenton reactions, leading to the generation of the toxic hydroxyl radical. Thus, intracellular iron levels must be tightly controlled both to satisfy physiological needs and to avoid toxic effects.

In Gram-negative bacteria, cellular iron homeostasis is achieved by the ferric uptake regulator (Fur), which is the master repressor for iron uptake system genes, including siderophore biosynthesis and transport genes (5). When iron is replete, Fur complexed with ferrous iron binds to Fur boxes (6) within the promoter regions of Fur-regulated genes and represses their transcription. In contrast, when iron is depleted, Fur, inactivated by the loss of an iron corepressor, is released from Fur boxes, so that Fur-regulated iron uptake system genes are transcribed.

Bacterial small RNAs (sRNAs) are noncoding antisense transcripts, usually smaller than 300 nucleotides, that are involved in up- and downregulation of translation through base pairing with their target mRNAs (7–9). Most sRNAs are induced or repressed under specific environmental conditions, such as iron limitation (10), oxidative stress (11, 12), and glucose starvation (13, 14), and often they participate in the global response of genes involved in environmental adaptation (7). In most cases, sRNAs exert their regulatory function by short base pairing with the 5′-untranslated regions (5′-UTRs) of target mRNAs. Typically, sRNA pairing masks the ribosomal binding site (RBS) and/or initiation codon of its target mRNA, preventing ribosome entry in the mRNA and inhibiting translation initiation (15). The untranslated target mRNA is degraded rapidly, usually due to the action of RNase E (16) or RNase III (17); however, there are some cases of translational repression without the change of mRNA stability. For example, sRNA Spot42 blocks the initiation of translation of galK in galETKM mRNA without significantly affecting the stability of galETKM mRNA (13). In addition to downregulation of target mRNA via RBS blocking by sRNA, there are several cases of translational inhibition of target mRNA by sRNA binding far upstream of the RBS (18). In other cases, some sRNAs base pair with the coding sequence of target mRNA (19, 20) or with the intergenic region of polycistronic target mRNA (21), and thereby these target mRNAs are decayed. In contrast, sRNAs can also activate target mRNAs, in many cases, via the anti-antisense mechanism; base pairing of sRNAs with the inhibitory structure of target mRNA, in which the RBS is sequestered, gives rise to remodeling of the mRNA structure, thereby unmasking the RBS and promoting translation (8). As another example of upregulation of target mRNA mediated by sRNAs, it was reported that an Escherichia coli sRNA, gadY, was on the opposite strand between gadX and gadW base pairs, with the 3′-UTR of gadX to stabilize this transcript and activate translation (22). On the other hand, base pairing between target mRNA and sRNA usually requires the assistance of Hfq (23), an RNA binding protein that forms a ring-shaped homohexamer of ∼11-kDa subunits (24), which was first identified in E. coli as a host factor essential for bacteriophage Qβ replication (25). Hfq is homologous to the eukaryotic and archaeal Sm and Sm-like proteins, which are engaged in various aspects of RNA metabolism, including splicing and mRNA decay (26). Hfq can bind RNA, preferably at single-stranded AU-rich sequences near stem-loop structures (13, 27), the poly(A) tracts of mRNAs (28), and the poly(U) tails of sRNAs (29–32). Thus, Hfq functions as an RNA chaperone by facilitating the imperfect base-pairing interactions between sRNAs and target mRNAs, thereby modulating translation efficiency and/or resulting in cleavage of both RNAs by RNase E (23, 33). Hfq binding to sRNAs is also known to protect sRNAs from degradation by RNase E through occluding the RNase E cleavage sites on sRNAs (34, 35).

RyhB, one of the well-studied sRNAs, was first identified in E. coli by Massé and Gottesman (10) as a Fur-regulated sRNA characteristically involved in the modulation of iron metabolism through the repression of iron-using and iron storage genes, such as sdhCDAB (encoding succinate dehydrogenase), acnA (aconitase) and fumA (fumarase) (tricarboxylic acid [TCA] cycle enzyme genes), sodB (Fe superoxide dismutase gene), and bfr and ftnA (iron storage genes) (10, 36), by base pairing with these target mRNAs. Likewise, RyhB homologs that negatively regulate almost the same genes as those in E. coli in response to iron starvation have been found in other Gram-negative bacteria, such as Vibrio cholerae (37–39), Shigella species (40), and Pseudomonas aeruginosa (41). In these cases, the resulting RyhB-mRNA duplexes usually are subject to subsequent degradation mediated by RNase E (42, 43), thereby reducing the level of iron-using and iron storage proteins to modulate intracellular iron usage.

V. parahaemolyticus is a halophilic Gram-negative bacterium that typically inhabits sea and brackish water, and it can cause diarrhea and gastroenteritis upon consumption of seafood contaminated with the bacterium (44, 45). It has been previously reported that V. parahaemolyticus is able to utilize some siderophores, including the cognate vibrioferrin (46), exogenous aerobactin (47), ferrichrome (48), and enterobactin (49), to acquire iron under iron-depleted conditions. The Fur-regulated gene cluster involved in the vibrioferrin-mediated iron acquisition system is composed of 2 divergent operons that contain pvsABCDE (pvsABDE, vibrioferrin biosynthesis genes; pvsC, vibrioferrin exporter gene) (Fig. 1A) and pvuA1-A2-BCDE (pvuA1 and pvuA2, outer membrane receptor genes for ferric vibrioferrin; pvuBCDE, ABC transport system genes for ferric vibrioferrin) (50–52). In this study, we found that the production of vibrioferrin greatly decreased in the V. parahaemolyticus ryhB deletion mutant relative to the wild-type parental strain. Furthermore, we provide evidence that the 5′-UTR within the pvsABCDE operon (here called pvsOp) possesses a potential RyhB base-pairing region, and that the formation of the RyhB-pvsOp 5′-UTR duplex with the help of Hfq stabilizes the pvsOp mRNA, leading to the enhanced production of vibrioferrin. To our knowledge, this is the first report that RyhB positively regulates a polycistronic mRNA involved in siderophore biosynthesis as a direct target.

Fig 1.

(A) Arrangement of pvsOp (VPA1658 to VPA1662) located on chromosome 2 of V. parahaemolyticus. The orientation of genes in pvsOp is shown by arrows. A closed box and a wavy arrow indicate a putative Fur box and pvsOp mRNA, respectively. The pvsABDE genes encode the vibrioferrin biosynthetic enzymes, and pvsC encodes the exporter of vibrioferrin (50, 51). (B) Nucleotide sequence of the pvsOp 5′-UTR with a potential base-pairing site with RyhB. The potential region for base pairing between RyhB and the pvsOp 5′-UTR was determined using the TargetRNA program. The probable ribosomal binding site (RBS) and the initiation codon of pvsA are indicated by an underline and by large letters, respectively. The transcription start site (+1) of pvsOp, determined by primer extension, is shown by a right-angled arrow. The numbers flanking the sequences of the pvsOp mRNA and RyhB indicate the nucleotide positions relative to the transcriptional start site (+1).

MATERIALS AND METHODS

Bacterial strains, plasmids, primers, and growth conditions.

The bacterial strains and plasmids used in this study are listed in Table 1. The PCR primers used in this study are listed in Table S1 in the supplemental material. E. coli strains were routinely grown in LB medium containing 0.5% NaCl; however, growth and maintenance of E. coli β2155 (53) required 0.5 mM 2,6-diaminopimelic acid. V. parahaemolyticus strains were incubated at 37°C in LB (iron-replete) medium containing 3% NaCl unless otherwise noted. To impose iron limitation on V. parahaemolyticus strains, they were usually grown in LB medium containing 25 μM ethylenediamine-di(o-hydroxyphenylacetic acid) (EDDA; Sigma-Aldrich); however, LB medium containing 200 μM 2,2′-dipyridyl (DPD; Wako Pure Chemical Industries, Osaka, Japan) instead of EDDA was used for rifampin chase experiments (see below), because V. parahaemolyticus RIMD2210633 (O3:K6), used as the wild-type parental strain in this study, was found to be sensitive to rifampin in iron-depleted LB plus DPD medium for some unknown reason, although approximately 70% of V. parahaemolyticus O3:K6 strains have been reported to be resistant to this drug (57). Antibiotics were used at the following concentrations: ampicillin, 100 μg/ml; chloramphenicol, 10 μg/ml; and tetracycline, 10 μg/ml.

Table 1.

Bacterial strains and plasmids used in this study

Strain or plasmid	Descriptiona	Reference or source
Strains
V. parahaemolyticus
RIMD2210633	Clinical isolate of serotype O3:K6; wild-type strain	56
VPD5	Deletion mutant in pvsB (vibrioferrin biosynthesis gene) of RIMD2210633; vibrioferrin deficient	52
ΔryhB	Deletion mutant in ryhB of RIMD2210633	This study
Δhfq	Deletion mutant in hfq of RIMD2210633	This study
pvsOpΔ86-104	Deletion mutant in the pvsOp86-104 region of RIMD2210633	This study
ΔryhB pvsOpΔ86-104	Double deletion mutant in the ryhB and pvsOp86-104 region of RIMD2210633	This study
pvsOp86-95comp	Complementary replacement mutant of the pvsOp86-95 region from RIMD2210633	This study
pvsOp86-94comp	Complementary replacement mutant of the pvsOp86-94 region from RIMD2210633	This study
pvsOp86-93comp	Complementary replacement mutant of the pvsOp86-93 region from RIMD2210633	This study
pvsOp86-92comp	Complementary replacement mutant of the pvsOp86-92 region from RIMD2210633	This study
pvsOp86-91comp	Complementary replacement mutant in the pvsOp86-91 region of RIMD2210633	This study
pvsOp86-90comp	Complementary replacement mutant of the pvsOp86-90 region from RIMD2210633	This study
pvsOp86-89comp	Complementary replacement mutant of the pvsOp86-89 region from RIMD2210633	This study
pvsOp86-88comp	Complementary replacement mutant of the pvsOp86-88 region from RIMD2210633	This study
pvsOp86-87comp	Complementary replacement mutant of the pvsOp86-87 region from RIMD2210633	This study
ryhB139-144comp	Complementary replacement mutant in the ryhB139-144 region of RIMD2210633	This study
pvsOp86-91comp ryhB139-144comp	Complementary replacement mutant in the pvsOp86-91 and ryhB139-144 regions of RIMD2210633	This study
E. coli
β2155	thrB1004 pro thi strA hsdS Δ(lacZ)ΔM15 [F′ Δ(lacZ)M15 lacIq traD36 proA+ proB+] ΔdapA::erm(Emr), pir::RP4; kan(Kmr) from SM10)	53
DH5α	F− endA1 glnV44 thi-1 recA1 relA1 gyrA96 deoR nupG Φ80dlacZΔM15 Δ(lacZYA-argF)U169, hsdR17(rK− mK+), λ−; host for cloning of pET-21b(+)-hfq	Promega
BL21(DE3)pLysS	F− ompT hsdSB(rB− mB−) gal dcm lon λ(DE3), pLysS (Cmr)	Invitrogen
Plasmids
pXAC623	Suicide vector derived from pKTN701 containing the sacB gene of Bacillus subtilis; Cmr	55
pRK415	Broad-host-range plasmid (∼10.5 kb); Tcr	54
pET-21b(+)	T7 expression plasmid; Apr	Merck Millipore
pXACΔryhB	pXAC623 containing 951-bp XbaI-XbaI fragment with 111-bp deletion in ryhB; Cmr	This study
pXACΔhfq	pXAC623 containing 1,683-bp XhoI/SalI-XhoI/SalI fragment with 167-bp deletion in hfq; Cmr	This study
pXACΔpvs86-104	pXAC623 containing 1,325-bp XhoI/SalI-XhoI/SalI fragment with 19-bp deletion in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-95comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 10-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-94comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 9-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-93comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 8-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-92comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 7-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-91comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 6-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-90comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 5-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-89comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 4-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-88comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 3-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACpvs86-87comp	pXAC623 containing 1,344-bp XhoI/SalI-XhoI/SalI fragment with 2-base complementary replacement in pvsOp 5′-UTR; Cmr	This study
pXACryhB139-144comp	pXAC623 containing 1,062-bp XbaI-XbaI fragment with 6-base complementary replacement in ryhB; Cmr	This study
pRK415-ryhB	pRK415 containing 1,062-bp XbaI-XbaI fragment with full-length ryhB; Tcr	This study
pRK415-hfq	pRK415 containing 1,750-bp EcoRI-EcoRI fragment with full-length hfq; Tcr	This study
pET21b-hfq	pET-21b(+) containing 263-bp NdeI-XhoI fragment with full-length hfq; Apr	This study

^aAp^r, ampicillin resistance; Cm^r, chloramphenicol resistance; Tc^r, tetracycline resistance.

Primer extension.

Stationary-phase cells of V. parahaemolyticus RIMD2210633 were inoculated into LB medium or LB plus EDDA medium at a final optical density at 600 nm (OD₆₀₀) of 0.005, and the cultures were then shaken at 37°C until they reached an OD₆₀₀ of 0.5. The total RNA was extracted with the RNeasy Protect bacterial Mini kit (Qiagen). Primer extension of ryhB was carried out using the VpryhB-PE primer (see Table S1 in the supplemental material), 5′ labeled with Texas Red, and an SQ5500 DNA sequencer (Hitachi, Tokyo, Japan) as previously described (49).

Construction of deletion and complementary replacement mutants.

The deletion and complementary replacement mutants (in which nucleotides within the pvsOp 5′-UTR and/or ryhB are replaced by the corresponding complementary nucleotides [see Fig. S1 in the supplemental material for sequence details]) listed in Table 1 were constructed by double-crossover allelic exchange using the R6K-ori suicide vector pXAC623 (55). DNA fragments with deletions in ryhB, hfq, and the pvsOp 5′-UTR and with complementary replacement in ryhB and the pvsOp 5′-UTR were prepared by PCR-driven overlap extension (58), as previously described (52). The PCR fragments were digested with appropriate restriction enzymes and cloned into pXAC623. Each resulting plasmid was transformed into E. coli β2155 and then mobilized into the relevant V. parahaemolyticus strains by filter mating. The chloramphenicol-resistant merodiploids thus obtained were spread on VDS-broth agar plates (1% polypeptone, 0.5% yeast extract, 30 mM NaCl, 55 mM KCl, 10% sucrose, and 2.5% agar) at 25°C for 30 h, and sucrose-resistant and chloramphenicol-sensitive colonies were selected. All mutants were verified by PCR and/or sequencing (data not shown).

Vibrioferrin production and growth assays.

Vibrioferrin production by V. parahaemolyticus was visually assessed by the chrome azurol S (CAS) agar plate assay and/or CAS liquid assay (59). The principle of the CAS assay is based on a color change of CAS-iron complex from blue (CAS-iron-hexadecyltrimethylammonium [HDTMA] complex) to orange (CAS-HDTMA complex) resulting from the removal of the bound iron from the blue dye by siderophores. For CAS plate assay, V. parahaemolyticus strains were first grown on LB plates for 16 to 24 h at 37°C, and single-colony cells were spotted with a sterilized toothpick onto CAS agar plates containing 2% NaCl and incubated for 48 to 72 h at 30°C. For CAS liquid assay, V. parahaemolyticus strains were precultured in Tris-buffered succinate (TS) medium (12.1 g of Tris, 3.7 g of KCl, 1.1 g of NH₄Cl, 0.272 g of KH₂PO₄, 0.15 g of CaCl₂·2H₂O, 0.142 g of Na₂SO₄, 0.1 g of MgCl₂ · 6H₂O, 5 g of disodium succinate per liter; pH 7.4) containing 2% NaCl and 1 μM FeCl₃ at 30°C for 24 h. The precultures were then diluted 1:50 in fresh TS medium containing 2% NaCl and 0.2 μM FeCl₃ and incubated at 30°C for 24 h. The cultures were centrifuged, and 0.5 ml of the clear supernatant was used for the CAS liquid assay. Growth assays of V. parahaemolyticus strains were performed in iron-depleted LB plus EDDA medium with a biophotorecorder (TVS062CA; Advantec Toyo, Tokyo, Japan). When required, the iron-depleted medium was supplemented with vibrioferrin (46) at a final concentration of 20 μM.

Purification of the V. parahaemolyticus Hfq protein.

To prepare the C-terminal His tag-fused Hfq, the expression vector pET-21b(+)-hfq was constructed by ligating the PCR product amplified with a set of primers, Vphfq-Nde-F and Vphfq-XhoI-R (see Table S1 in the supplemental material), into the NdeI/XhoI sites of pET-21b(+). The primers were designed based on the sequence of the flanking region of hfq (VP2817), and E. coli DH5α was used as a host for the cloning of pET-21b(+)-hfq. E. coli BL21(DE3)pLysS/pET-21b(+)-hfq, used for Hfq overexpression, was grown in iron-replete medium with ampicillin and chloramphenicol at 37°C to an OD₆₀₀ of 0.3, and Hfq expression then was induced by the addition of isopropyl thiogalactoside at a final concentration of 0.5 mM to the medium, followed by additional incubation for 1.5 h. Cells were harvested by centrifugation, washed with STE buffer (100 mM NaCl, 10 mM Tris-HCl, 1 mM EDTA, pH 8), and disrupted by sonication in buffer A (50 mM NaH₂PO₄, 300 mM NaCl, pH 8) containing 10 mM imidazole. After centrifugation, the lysate was treated with 10 μg/ml RNase A at room temperature for 60 min, boiled at 80°C for 10 min, and then centrifuged again. The supernatant obtained was loaded onto a HisTrap HP column (GE Healthcare, Buckinghamshire, United Kingdom), washed with buffer A containing 20 mM imidazole, and eluted with buffer A containing 250 mM imidazole. Pooled fractions containing Hfq were concentrated and desalted by buffer exchange into buffer B (10 mM Tris-HCl, 50 mM NaCl, 50 mM KCl, 1 mM EDTA, 10% glycerol, pH 8) using a Vivaspin 20-3K (Sartorius, Gottingen, Germany). The concentration of the protein was assayed by the Lowry method. The concentrations of His tag-fused Hfq are indicated with respect to its hexameric form throughout this work.

In vitro RNA transcription.

RyhB and RyhB^139-144comp RNAs were synthesized by in vitro transcription with T7 RNA polymerase (Roche). For this purpose, the PCR product used as the template of RNA synthesis was generated with a pair of primers, T7-VpryhB-F and T7-VpryhB-R (see Table S1 in the supplemental material), from genomic DNAs isolated from the wild-type strain, RIMD2210633, and the ryhB^139-144comp complementary replacement mutant. Transcription was performed in transcription buffer (40 mM Tris-HCl, 6 mM MgCl₂, 10 mM dithiothreitol, 2 mM spermidine, pH 8) containing 1 mM nucleoside triphosphate (NTP) mix, 40 U RNasin (Promega), 20 U T7 RNA polymerase, and 100 ng of the DNA template amplified with PCR. After incubation for 4 h at 37°C, the reaction mixture was treated with 1 U RQ1 DNase (Promega) and purified with a ProbeQuant G50 microcolumn (GE Healthcare) followed by ethanol precipitation. Both the fluorescein-labeled pvsOp 5′-UTR and pvsOp^86-91comp 5′-UTR were transcribed in vitro in the same way, except that a fluorescein RNA labeling mix (Roche), PCR products generated from the primer pair T7-pvsOp-F and T7-pvsOp-R (see Table S1), and the genomic DNAs extracted from RIMD2210633 and the pvsOp^86-91comp complementary replacement mutant were used.

EMSA of the interaction between RyhB and pvsOp 5′-UTR.

Electrophoretic mobility shift assay (EMSA) was performed with the fluorescein-labeled pvsOp 5′-UTR or pvsOp^86-91comp 5′-UTR as a probe. The fluorescein-labeled transcript and unlabeled RyhB transcript, at the indicated concentrations, and 6.6 μM yeast tRNA (Ambion) were heat denatured at 90°C for 2 min and then slowly cooled to 37°C. The resulting solutions were mixed in EMSA buffer (10 mM Tris-HCl, 5 mM Mg acetate, 100 mM NH₄Cl, 0.5 mM dithiothreitol, pH 7.5) with or without purified Hfq at the indicated concentrations, and the EMSA reaction mix (10 μl in total) was incubated for 10 min at 37°C and cooled on ice for 1 min. Two microliters of 50% glycerol was then added to each sample, which was loaded onto a 5% native polyacrylamide gel in Tris-borate-EDTA buffer (90 mM Tris, 90 mM boric acid, 2 mM EDTA, pH 8.3) at 4°C to separate the complexes. Gel images were analyzed with a LAS-3000 gel imager (Fujifilm, Tokyo, Japan) using an excitation wavelength of 460 nm and an emission band-pass filter of 515 nm.

RT-qPCR analysis.

Stationary-phase V. parahaemolyticus cells were inoculated into LB medium and LB plus EDDA medium at a final OD₆₀₀ of ∼0.01, and the cultures were then shaken at 37°C until they reached an OD₆₀₀ of 0.3 to 0.6. The cultures obtained were treated with the RNAprotect bacterial reagent (Qiagen) according to the manufacturer's protocol. Total RNA samples were isolated with the TriPure isolation reagent (Roche, Basel, Switzerland) and treated with RNase-free DNase I (Ambion), and a 0.5-μg aliquot of RNA was reverse transcribed (RT) by using ReverTra Ace reverse transcriptase (Toyobo, Osaka, Japan) with a random hexamer primer (TaKaRa, Shiga, Japan). Quantitative PCR (qPCR) analysis was performed using the specific primer pairs VppvsA-qF and VppvsA-qR (for pvsA) and VppvsE-qF and VppvsE-qR (for pvsE) (see Table S1 in the supplemental material) with the Thunderbird SYBR qPCR mix (Toyobo) in a Chromo4 real-time PCR detection system (Bio-Rad). The relative mRNA expression levels were determined by the comparative threshold cycle method using the 16S rRNA expression level as an internal control. RT-qPCR primers for 16S rRNA are listed in Table S1 in the supplemental material. The linearity of the qPCRs was ensured by sequentially diluting cDNA samples to give the log values accompanied by proportionally increased values of threshold cycles.

Determination of stability of pvsOp mRNA by rifampin chase experiment.

Stationary-phase cells of V. parahaemolyticus strains were inoculated into LB medium at a final OD₆₀₀ of 0.005 and incubated at 37°C until the cultures reached an OD₆₀₀ of 0.1 to 0.2. The culture was separated into 2 aliquots. One was fortified with DPD at 200 μM and the other was not, and both were further incubated at 37°C until they reached an OD₆₀₀ of 0.3 to 0.6 for use in RT-qPCR. For the rifampin chase experiments, these cultures were further incubated with rifampin at 250 μg/ml at room temperature and recovered at time points of 0, 1, 2, 4, 6, 8, and 10 min for RT-qPCR. RT-qPCR analysis was performed for pvsA using the specific primer pair VppvsA-qF and VppvsA-qR (see Table S1 in the supplemental material).

RESULTS

Location, alignment, and iron-regulated expression of V. parahaemolyticus ryhB.

Mey et al. (38) first indicated that in Vibrio species, V. cholerae RyhB has diverse functions, ranging from iron homeostasis to the regulation of biofilm formation. Mey et al. (38) also identified a putative V. parahaemolyticus ryhB in a BLAST search of the V. parahaemolyticus genome sequence but did not carry out any detailed studies on its functions, although V. parahaemolyticus RyhB was also predicted to act on various target mRNAs demonstrated in V. cholerae. V. parahaemolyticus ryhB is located on chromosome 1 between VP0105 and VP0107, overlapping VP0106 and positioned in the opposite orientation (Fig. 2A) (the functions of the 3 other open reading frames are unknown), and possesses a putative Fur box in the promoter region (Fig. 2B), like the ryhB genes in V. cholerae (38) and E. coli (10). The transcriptional start site for V. parahaemolyticus ryhB was determined by primer extension with total RNAs from cells grown in iron-replete LB or iron-depleted LB plus EDDA medium. An extended product was detected only when total RNA from cells grown in iron-depleted medium was used as a template, and the transcriptional start site was also determined as shown in Fig. 2C. This is consistent with the presence of a putative Fur box, accounting for the iron-regulated expression of ryhB. Incidentally, the transcriptional start site for V. parahaemolyticus ryhB corresponded with that for V. cholerae ryhB, as confirmed by the 5′ rapid amplification of cDNA ends (RACE) method (38). In addition, because no lower-molecular-weight species were detected by primer extension, it is unlikely that the RyhB transcript is further processed.

Fig 2.

Location and alignment of V. parahaemolyticus ryhB and determination of its transcriptional start site. (A) Arrangement of the ryhB gene and its flanking region in V. parahaemolyticus. Genes are represented by arrows facing the direction of transcription. ryhB (black arrow) overlaps with VP0106. (B) Sequence features of possible functional significance in ryhB genes. The ryhB sequences of V. parahaemolyticus (Vp), V. cholerae (Vc), and E. coli (Ec) are aligned. Asterisks and dashes denote identical bases between Vp and Vc and among the 3 species, respectively. The putative Fur boxes (14 matches of 19 nucleotides in the E. coli consensus Fur box) (6) are boxed, the putative −10 and −35 promoter elements are overlined, the transcriptional start site (+1) of ryhB is shown by a right-angled arrow, and the rho-independent terminator is indicated by converging arrows. The highly conserved region among the ryhB sequences of 3 species is boxed with a dashed line. Potential AU-rich and poly(U) tail sequences as the binding sites of V. parahaemolyticus Hfq are denoted with heavy overlines. The numbers flanking the sequences of pvsOp mRNA and RyhB indicate nucleotide positions relative to the transcriptional start site (+1). (C) Determination of the transcriptional start site of V. parahaemolyticus ryhB by primer extension (PE). Reverse transcription was performed with the VpryhB-PE primer for total RNA isolated from the wild-type strain RIMD2210633 grown in iron-replete LB medium or iron-depleted LB plus EDDA medium. The same primer was also used to generate a sequence ladder (A, C, G, T). The transcriptional start site is labeled +1 (as for panel B).

Involvement of RyhB and Hfq in the normal production of vibrioferrin.

In recent years, E. coli RyhB has also been identified to act as a positive regulator of the production of its cognate siderophore, enterobactin, under iron-depleted conditions (60, 61). The regulation of gene expression by sRNAs, including RyhB, generally requires Hfq for base pairing with their target mRNAs (32). Therefore, to clarify whether RyhB and Hfq participate in the normal production of the cognate siderophore vibrioferrin, ryhB and hfq deletion mutants were constructed for use in CAS assay. While vibrioferrin production in the ΔryhB mutant was greatly diminished compared to that of wild-type strain RIMD2210633, the Δhfq mutant revealed a complete lack of vibrioferrin production, as observed for the vibrioferrin-deficient mutant VPD5 (52) (Fig. 3A). Moreover, complementation of the ΔryhB and Δhfq mutants with the full-length ryhB and hfq genes, respectively, restored vibrioferrin production at levels similar to that of the wild-type strain (Fig. 3A). The growth of the ΔryhB and Δhfq mutants under iron-depleted conditions was suppressed as well as that of the vibrioferrin-deficient VPD5 mutant, whereas the ΔryhB/pRK415-ryhB and Δhfq/pRK415-hfq complementing strains displayed growth similar to that of the wild-type strain (Fig. 3B). Moreover, the addition of vibrioferrin at 20 μM to the same medium partly compensated for the growth in a ΔryhB or Δhfq mutant background (Fig. 3B). Taken together, these results suggest that both RyhB and Hfq are at least involved in the normal production of vibrioferrin under iron-depleted conditions. In addition, because Hfq can protect RyhB from degradation by RNase E (16, 35), the defect in vibrioferrin production in the Δhfq mutant might be due largely to the degradation of RyhB.

Fig 3.

(A) CAS plate assay phenotypes of the wild-type parental strain V. parahaemolyticus RIMD2210633, strain VPD5, ΔryhB and Δhfq mutant strains, and complementing strains. A faintly orange halo in the ΔryhB strain and a blue color in VPD5 and Δhfq strains represent poor and no production of vibrioferrin, respectively. (B) Growth assay of RIMD2210633, VPD5, and ΔryhB, Δhfq, ΔryhB/pRK415-ryhB, and Δhfq/pRK415-hfq mutant strains. Each strain was grown at 37°C with shaking at 70 rpm in the iron-depleted LB plus EDDA medium, and cultures were monitored by measuring the OD₆₀₀ every 3 h for 24 h. Data are shown as means ± standard deviations (SD) from 3 separate experiments.

Detection of Hfq-mediated RyhB-pvsOp 5′-UTR base pairing by EMSA.

In general, many sRNAs modulate gene expression via base pairing with their target mRNAs. Therefore, to pursue the possibility that V. parahaemolyticus RyhB directly exerts a positive regulatory effect on pvsOp, which is required for the biosynthesis and export of vibrioferrin, we performed bioinformatics analysis of the pvsOp 5′-UTR using the TargetRNA program (http://snowwhite.wellesley.edu/targetRNA/index2.html) (62) to identify a potential region within the pvsOp 5′-UTR, with the result that a potential site for base pairing with RyhB (a stretch of 27 nucleotides complementary to RyhB) was found within the pvsOp 5′-UTR (Fig. 1B and 2B). To test whether the pvsOp 5′-UTR can actually form an RNA-RNA duplex with RyhB, we performed EMSA for RyhB in the presence of Hfq using fluorescein-labeled pvsOp 5′-UTR as a probe. As shown in Fig. 4A, panel a, the complex band formed between the pvsOp 5′-UTR and RyhB was gradually increased in intensity by RyhB in a concentration-dependent manner, even in the absence of Hfq, suggesting that the pvsOp 5′-UTR can bind to RyhB without the help of Hfq. Furthermore, because Hfq was found to be associated with vibrioferrin production at a normal level (Fig. 3A), His-tagged Hfq was also used to confirm its function by EMSA. As shown in Fig. 4A, panels b to d, Hfq bound to the pvsOp 5′-UTR in a concentration-dependent manner to be shifted as the pvsOp 5′-UTR-RyhB-Hfq band at increased levels, indicating that Hfq can bind to the RyhB-pvsOp 5′-UTR duplex and, as a consequence, promote the formation of the pvsOp 5′-UTR-RyhB-Hfq complex. When Hfq was used at a higher concentration (100 nM), an additional band that migrated more slowly than the pvsOp 5′-UTR-RyhB-Hfq complex was also detected by EMSA (Fig. 4A, panel d, asterisk). This band presumably is due to binding of a second Hfq to another affinity site on the pvsOp 5′-UTR; therefore, it was judged as a pvsOp 5′-UTR:RyhB:Hfq complex with 1:1:2 stoichiometry [here called pvsOp 5′-UTR-RyhB-(Hfq)₂]. Hfq is known to bind preferentially to AU-rich single-strand sequences in the proximity of double-stranded regions (13, 27) and to poly(A) tracts (28) in sRNAs. Consistent with this, possible regions targeted by Hfq were found in the largest loop of the folded pvsOp 5′-UTR predicted by the mfold Web server (http://mfold.rna.albany.edu/?q=mfold/RNA-Folding-Form) (63) (see Fig. S2 in the supplemental material). EMSA also showed that, like E. coli RyhB, V. parahaemolyticus RyhB can bind to Hfq (data not shown), which may be supported by the presence of a potential AU-rich sequence for Hfq binding in V. parahaemolyticus RyhB (Fig. 2B). In addition, a poly(U) tail of the rho-independent terminator that has been shown to function as an Hfq binding site in sRNAs (29–32) was found in V. parahaemolyticus RyhB (Fig. 2B). The EMSA bands were analyzed by gel densitometry to calculate the percentage of the whole probe shifted by binding with RyhB (Fig. 4B). The percentages of the probe shifted as ternary complexes [pvsOp 5′-UTR-RyhB-Hfq and pvsOp 5′-UTR-RyhB-(Hfq)₂] in the presence of Hfq were greatly increased compared to that of the probe shifted as a binary complex (pvsOp 5′-UTR-RyhB) in the absence of Hfq and were gradually increased as the Hfq concentration was increased. These results indicate that Hfq facilitates the base-pairing interaction between RyhB and the pvsOp 5′-UTR.

Fig 4.

Involvement of Hfq in base pairing between RyhB and pvsOp 5′-UTR. (A) Complex formation of the pvsOp 5′-UTR with RyhB and Hfq was analyzed by EMSA, which was performed using 50 nM fluorescein-labeled pvsOp 5′-UTR as a probe in the presence of Hfq hexamer at concentrations of 0, 50, 75, and 100 nM and RyhB at concentrations of 0, 5, 10, 25, 50, and 100 nM. The probe-RyhB, probe-RyhB-Hfq, and probe-RyhB-(Hfq)₂ complexes formed are indicated with white and black arrowheads and an asterisk, respectively. (B) Based on the results shown in panel A, the sum of the probe shifted as probe-RyhB, probe-RyhB-Hfq, and probe-RyhB-(Hfq)₂, except for probe-Hfq, was determined by gel densitometry and is shown as a percent relative to the total amount of probe used.

Involvement of the pvsOp 5′-UTR in the production of vibrioferrin.

To investigate whether the potential RyhB pairing region of the pvsOp 5′-UTR is engaged in the regulation of vibrioferrin production, the regions ranging from nucleotide positions +86 to +104 relative to the transcription start site (+1) of pvsOp were deleted from the wild-type strain and the ΔryhB mutant to construct the pvsOp^Δ86-104 and ΔryhB pvsOp^Δ86-104 mutant strains, respectively (Table 1; also see Fig. S1 in the supplemental material). The region between nucleotide positions +86 and +104 is included in a potential RyhB binding region (Fig. 1B), and the pvsOp^Δ86-104 5′-UTR was unable to form a duplex with RyhB, as judged by EMSA (data not shown). Interestingly, RT-qPCR analysis of pvsA and pvsE showed that, when grown in iron-depleted LB plus EDDA medium, the pvsOp^Δ86-104 mutant, in which the 5′-UTR was unable to bind to RyhB, expressed both pvsA and pvsE mRNAs at levels similar to those in the wild-type strain; furthermore, the ΔryhB pvsOp^Δ86-104 double mutant restored the reduced expression of both pvsA and pvsE observed for the ΔryhB mutant to an extent equal to that of the wild-type strain (Fig. 5A). In accordance with these observations by RT-qPCR analysis, the pvsOp^Δ86-104 and ΔryhB pvsOp^Δ86-104 deletion mutants (see Fig. S1), as well as the wild-type strain and the complementing ΔryhB/pRK415-ryhB strain, produced vibrioferrin at normal levels, as judged by CAS plate assays (Fig. 5B). Accordingly, all of these strains were able to grow in iron-depleted LB plus EDDA medium, in contrast to VPD5 and ΔryhB strains, both of which are deficient in vibrioferrin (Fig. 5C). Taken together, these results suggest that the RyhB binding region within the pvsOp 5′-UTR is involved in reduction of the pvsOp mRNA level when RyhB is absent, and that deletion of the RyhB binding region from the pvsOp 5′-UTR leads to normal levels of pvsOp mRNA regardless of the presence or absence of RyhB, thereby promoting the production of vibrioferrin. Moreover, because RT-qPCR data showed a similar tendency for relative expression levels of both pvsA and pvsE (Fig. 5A), RyhB was thought to affect the level of the entire pvsOp mRNA.

Fig 5.

(A) Effects of deletions in ryhB and the probable RyhB pairing region within the pvsOp 5′-UTR on pvsOp transcript levels. The relative mRNA levels of pvsA and pvsE within pvsOp were determined by RT-qPCR. Total RNA was extracted from RIMD2210633 and the ΔryhB, pvsOp^Δ86-104, and ΔryhB pvsOp^Δ86-104 strains grown in iron-replete LB and iron-depleted LB plus DPD media. Data are shown as means ± SD from 3 separate experiments. (B) CAS plate assay phenotypes of the ΔryhB mutant, the ΔryhB strain complemented with the native ryhB, and the pvsOp^Δ86-104 and ΔryhB pvsOp^Δ86-104 mutants. Assay conditions were the same as those described for Fig. 3A. The RIMD2210633 strain and the vibrioferrin-deficient mutant VPD5 were included as positive and negative controls. (C) Growth assay of the RIMD2210633 wild-type strain and the VPD5, ΔryhB, ΔryhB/pRK415-ryhB, pvsOp^Δ86-104, and ΔryhB pvsOp^Δ86-104 mutant strains. Growth conditions are the same as those described in the legend to Fig. 3B. Data are shown as means ± SD from 3 separate experiments.

Decrease in stability of pvsOp mRNA in the ΔryhB and Δhfq mutants.

Each of the ΔryhB and Δhfq mutants exhibited a significant decrease in vibrioferrin production (Fig. 3), and the pvsOp 5′-UTR was able to bind to RyhB and Hfq (Fig. 4). Furthermore, the ΔryhB mutant exhibited a decreased level of pvsOp mRNA compared to that of its parental strain, RIMD2210633 (Fig. 5A). Therefore, we assumed that binding of RyhB and Hfq to the pvsOp 5′-UTR contributes to the stabilization of pvsOp mRNA. This assumption was verified by measuring the time-dependent decay of pvsOp mRNA after rifampin exposure using RT-qPCR with a primer pair, VppvsA-qF and VppvsA-qR (see Table S1 in the supplemental material), that is specific to pvsA. The pvsOp transcripts in the ΔryhB and Δhfq mutants induced in iron-depleted LB plus DPD medium were degraded more rapidly after rifampin exposure than the parental strain RIMD2210633 (Fig. 6A, panels a to c). Based on the data presented in Fig. 6A, the mean percentages of pvsOp transcript levels relative to that at 0 min after rifampin exposure were plotted to construct decay curves to calculate the half-life of pvsOp mRNA (Fig. 6B). This approach showed that pvsOp mRNA had a half-life of ∼3.0 min in RIMD2210633 and half-lives of less than 1 min in both the ΔryhB and Δhfq mutants. Thus, both RyhB and Hfq exert a stabilization effect on pvsOp mRNA at the posttranscriptional level. It has been reported that the E. coli shiA mRNA, which is positively regulated by RyhB, has a half-life of 2 min and less than 1 min in the presence and absence of RyhB, respectively (60). These half-lives are comparable to those observed for pvsOp mRNA.

Fig 6.

Analysis of the stability of pvsOp transcripts by rifampin chase experiments. (A) The relative expression levels of pvsOp transcripts after rifampin treatment were determined by RT-qPCR. Rifampin (250 μg/ml) was added to V. parahaemolyticus cultures grown in the iron-depleted LB plus DPD medium when the OD₆₀₀ reached 0.6, and total RNAs were isolated from cells exposed to rifampin for 0, 1, 2, 4, 6, 8, and 10 min. Data are shown as means ± SD from 3 to 5 separate experiments. (B) Mean percentages of the remaining pvsOp mRNA levels relative to the 0-min time point are presented.

Confirmation of base pairing between RyhB and pvsOp 5′-UTR.

To confirm the involvement of RyhB-pvsOp 5′-UTR base pairing in the normal production of vibrioferrin, we constructed a complementary replacement mutant of the pvsOp 5′-UTR that neither binds to RyhB nor produces vibrioferrin at a normal level (i.e., a mutant having a phenotype similar to that of the ΔryhB strain). Among 9 replacement mutants generated, only pvsOp^86-91comp, in which the original TGGAAG sequence between nucleotide positions +86 and +91 in pvsOp was replaced by ACCTTC (see Fig. S1 in the supplemental material for details), exhibited negative phenotypes in CAS plate and liquid assays, as was seen with the ΔryhB mutant (Fig. 7A), and the 5′-UTR of the pvsOp^86-91comp mutant did not bind to RyhB, as judged by EMSA (Fig. 7B). Moreover, the original CTTCCA sequence between nucleotide positions +139 and +144 in RyhB, which was expected to base pair with the region ranging from nucleotide positions +86 to +91 in pvsOp, was replaced by GAAGGT, yielding the ryhB^139-144comp mutant (see Fig. S1). The ryhB^139-144comp mutant exhibited negative phenotypes in CAS plate and liquid assays (Fig. 7A), and RyhB^139-144comp transcript was almost unable to form a duplex with the pvsOp 5′-UTR, as judged by EMSA (Fig. 7B). Finally, we constructed a pvsOp^86-91comp ryhB^139-144comp double complementary replacement mutant in which pvsOp^86-91comp 5′-UTR and RyhB^139-144comp transcripts could base pair (see Fig. S1). As expected, the pvsOp^86-91comp ryhB^139-144comp double mutant restored vibrioferrin production at a level similar to that observed for RIMD2210633, as determined by CAS plate and liquid assays (Fig. 7A). Consistent with this, EMSA revealed that the RyhB^139-144comp transcript was able to bind to the pvsOp^86-91comp 5′-UTR (Fig. 7B). To test the stability of pvsOp, rifampin chase experiments followed by RT-qPCR were carried out for the complementary replacement mutants. Similar to the ΔryhB and Δhfq mutants (Fig. 6A, panels b and c), pvsOp transcripts in the pvsOp^86-91comp and ryhB^139-144comp mutants were rapidly degraded (Fig. 6A, panels d and e), with half-lives of around 1.5 min and less than 1 min, respectively (Fig. 6B). In contrast, the pvsOp transcript in the pvsOp^86-91comp ryhB^139-144comp mutant was as stable as that in RIMD2210633 (Fig. 6A, panel f), with a half-life of around 4 min (Fig. 6B). Taken together, these results demonstrate that RyhB-pvsOp 5′-UTR base pairing leads to the stabilization of pvsOp mRNA, thereby promoting vibrioferrin production. In addition, it has been recognized that some sRNAs are able to bind to the 5′-UTRs of their target mRNAs to impede the formation of inhibitory structures, so that translation of the target mRNAs is activated (reviewed in reference 8). However, the potential secondary structure of the pvsOp 5′-UTR that was predicted by the mfold Web server appeared incapable of forming an inhibitory structure by masking the RBS and translation initiation codon (see Fig. S2).

Fig 7.

Effect of base pairing between RyhB and the pvsOp 5′-UTR on vibrioferrin production. (A) Abilities of RIMD2210633, ΔryhB, pvsOp^86-91comp, ryhB^139-144comp, and pvsOp^86-91comp ryhB^139-144comp strains to produce vibrioferrin were assessed using CAS plate and CAS liquid assays. (B) EMSA was performed using the fluorescein-labeled pvsOp (normal) and pvsOp^86-91comp (86-91comp) 5′-UTR transcripts at 50 nM as the probes in the presence of Hfq (as hexamer), together with RyhB or RyhB^139-144comp at the indicated concentrations.

DISCUSSION

It is generally known that target mRNAs negatively regulated by sRNAs, including RyhB, form secondary structures in their 5′-UTRs to occlude RBSs and/or initiation codons, thereby impairing translational initiation and destabilizing target mRNAs (64). However, positive regulation of target mRNAs by sRNAs has been described for some sRNAs, such as DsrA (65, 66), RprA (67), and RNAIII (68). Recently, promotion of the stability of target mRNAs by sRNAs to activate the expression of virulence genes has been reviewed (8).

There have been 2 reports so far on the involvement of E. coli RyhB in promoting the synthesis of the siderophore enterobactin, which is assembled from the precursors 2,3-dihydroxybenzoic acid (DHBA) and serine. First, RyhB posttranscriptionally activates the expression of the shiA permease gene through direct positive regulation to effectively obtain shikimate, one of the precursors for the biosynthesis of DHBA, thereby increasing enterobactin production (60). In this case, the region for the potential base pairing with RyhB was detected in the 5′-UTR of shiA mRNA. Second, RyhB is required for the normal expression of entCEBAH mRNA, a transcript encoding DHBA, so that DHBA is sufficiently synthesized to allow the enhanced production of enterobactin (61). In addition, RyhB also contributes to retain serine for biosynthesis of enterobactin through the depression of serine catabolism mediated by CysE (serine O-acetyltransferase) by blocking the translation of cysE (61). Although evidence for the translation block of cysE by base pairing between RyhB and cysE mRNA has been provided, a region necessary for base pairing with RyhB has not been identified in the 5′-UTR of entCEBAH mRNA. On the other hand, V. parahaemolyticus Hfq (VP2817) has been reported to negatively regulate the production of thermostable direct hemolysin (69) and to reduce its antioxidative ability (70). However, to our knowledge, the involvement of RyhB and Hfq in siderophore production and transport has not been reported in this species.

Our present study provides evidence that V. parahaemolyticus RyhB and Hfq upregulate production of the cognate siderophore vibrioferrin. The CAS assays and growth assays showed that the ΔryhB and Δhfq mutants repressed the production of vibrioferrin and growth in iron-depleted LB plus EDDA medium (Fig. 3); however, the growth of those mutants was not fully restored by the addition of vibrioferrin at 20 μM (Fig. 3B), the concentration sufficient to restore the growth of the vibrioferrin-deficient mutant VPD5 (52). This might be explained by RyhB also being involved in positive regulation of the uptake and intracellular utilization systems for ferric vibrioferrin. In fact, the ΔryhB mutant showed approximately 40% reduction in expression of the vibrioferrin receptor gene pvuA1, which is the first gene in the ferric vibrioferrin transport operon pvuA1-pvuA2-pvuBCDE (52), compared to that of the parental strain RIMD2210633 (data not shown). In this context, it is interesting that V. cholerae RyhB has been reported to positively regulate expression of some genes involved in iron uptake (38).

It has been indicated that V. cholerae RyhB regulates the genes engaged in motility and chemotaxis, in addition to various genes encoding iron-containing proteins, such as sodB and fumA (38). However, the V. cholerae ΔryhB mutant did not impair growth in iron-depleted medium, and V. cholerae ryhB did not influence the expression of genes encoding its own siderophore, vibriobactin (37, 38). In addition, a BLASTN search indicated that no pvsOp 5′-UTR homolog was present in other Vibrio species, except for V. alginolyticus, which can produce vibrioferrin (71).

Table 2 summarizes the functional characteristics of the mutants constructed from the V. parahaemolyticus RIMD2210633 wild-type strain in this study. The deletions in ryhB and hfq were complemented by the corresponding genes, indicating at least that both genes are involved in the normal production of vibrioferrin. The experimental results shown in Fig. 3 and 4 indicate that the base pairing between the pvsOp 5′-UTR and RyhB is a requisite for the normal production of vibrioferrin. Interestingly, however, the pvsOp^Δ86-104 and ΔryhB pvsOp^Δ86-104 mutants showed a vibrioferrin-positive phenotype, although these deletion mutants would be unable to base pair between the pvsOp 5′-UTR and RyhB. One explanation for this is that the RyhB binding region, including the nucleotide positions 86 to 104, in the pvsOp 5′-UTR is important for the formation of a secondary structure of the pvsOp 5′-UTR involved in rapid degradation of pvsOp mRNA in the absence of RyhB, so that the pvsOp^Δ86-104 transcript, in which the RyhB binding region is deleted, is stable enough to produce vibrioferrin in the absence of RyhB. In addition, the experimental results obtained for the pvsOp^86-91comp, ryhB^139-144comp, and pvsOp^86-91comp ryhB^139-144comp mutants indicate that base pairing between the pvsOp 5′-UTR and RyhB is most important for the normal production of vibrioferrin. Meanwhile, 8 other complementary replacement mutants constructed, i.e., the pvsOp^86-95comp, pvsOp^86-94comp, pvsOp^86-93comp, pvsOp^86-92comp, pvsOp^86-90comp, pvsOp^86-89comp, pvsOp^86-88comp, and pvsOp^86-87comp mutants, like the pvsOp^Δ86-104 mutants, also produced an abundant vibrioferrin, even though RyhB could not bind or poorly bound to their pvsOp 5′-UTR (see Fig. S3 in the supplemental material). This may be due to the failure of these 8 mutants in constructing a secondary stem-loop structure within the pvsOp 5′-UTR that is responsible for rapid degradation of pvsOp mRNA. In contrast, the attenuation of vibrioferrin production observed for the pvsOp^86-91comp mutant may be explained by construction of a secondary structure within the pvsOp 5′-UTR of this mutant that is highly susceptible to rapid degradation of pvsOp mRNA.

Table 2.

Functional characteristics of the mutant and complementing strains constructed in this study

Mutant strain	Presence or absence ofa:
	Production of vibrioferrin	Base pairing between the pvsOp 5′-UTR and RyhB	Rapid degradation of pvsOp mRNAb
ΔryhB	−	−	+
ΔryhB/pRK415-ryhB	+	+	−
Δhfq	−	−	+
Δhfq/pRK415-hfq	+	+	−
pvsOpΔ86-104	+	−	−
ΔryhB pvsOpΔ86-104	+	−	−
pvsOp86-91comp	−	−	+
ryhB139-144comp	−	−	+
pvsOp86-91comp ryhB139-144comp	+	+	−

^aThe plus and minus symbols represent the presence and absence, respectively, of the functional characteristic indicated.

^bPresumed on the basis of the experimental results.

In conclusion, we showed that V. parahaemolyticus RyhB positively regulates the production of vibrioferrin through the stabilization of polycistronic pvsOp mRNA involved in its biosynthesis, namely, pvsOp mRNA transcribed under iron-depleted conditions would have a secondary structure which participates in the rapid degradation of pvsOp mRNA unless RyhB is present. Moreover, it is possible that V. parahaemolyticus RyhB activates translation of pvsOp mRNA by an anti-antisense mechanism, like translation activation of shiA mRNA by E. coli RyhB (60), although the secondary structure of the pvsOp 5′-UTR predicted by the mfold program suggested an unmasked RBS of pvsOp mRNA (see Fig. S2 in the supplemental material). Therefore, it could not be ruled out that the stabilization of pvsOp mRNA is also due to disruption of the inhibitory structure of the pvsOp 5′-UTR, leading to translation activation of pvsOp mRNA, which can protect it from rapid degradation (72). This must be clarified by further studies. The physiological role of pvsOp mRNA degradation in the absence of RyhB may be to rapidly respond to iron-replete conditions that may otherwise increase the intracellular iron pool to noxious levels, because the expression of RyhB is simultaneously repressed under iron-replete conditions. The posttranscriptional regulation of V. parahaemolyticus pvsOp mRNA is the first example of positive regulation by RyhB of translation of the polycistronic transcript involved in siderophore biosynthesis as its direct target. Further studies should clarify the molecular mechanism by which pvsOp mRNA is stabilized by RyhB and Hfq and is rapidly degraded in the absence of RyhB.