This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Boardman, B. K.
Right arrow Articles by Fullner Satchell, K. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Boardman, B. K.
Right arrow Articles by Fullner Satchell, K. J.

 Previous Article  |  Next Article 

Journal of Bacteriology, March 2007, p. 1827-1835, Vol. 189, No. 5
0021-9193/07/$08.00+0     doi:10.1128/JB.01766-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Growth Phase Regulation of Vibrio cholerae RTX Toxin Export{triangledown}

Bethany Kay Boardman, Brian M. Meehan,{dagger} and Karla J. Fullner Satchell*

Department of Microbiology-Immunology, Northwestern University Feinberg School of Medicine, Tarry 3-713, 303 E. Chicago Avenue, Chicago, Illinois 60611

Received 20 November 2006/ Accepted 13 December 2006


arrow
ABSTRACT
 
Vibrio cholerae, the causative agent of the severe diarrheal disease cholera, secretes several "accessory" toxins, including RTX toxin, which causes the cross-linking of the actin cytoskeleton. RTX toxin is exported to the extracellular milieu by an atypical type I secretion system (T1SS), and we previously noted that RTX-associated activity is detectable only in supernatant fluids from log phase cultures. Here, we investigate the mechanisms for regulating RTX toxin activity in supernatant fluids. We find that exported proteases are capable of destroying RTX activity and may therefore play a role in the growth phase regulation of toxin activity. We determined that the absence of RTX toxin in stationary-phase culture supernatant fluids is also due to a lack of toxin secretion and not attributable to solely proteolytic degradation. We ascertained that the T1SS apparatus is regulated at the transcriptional level by growth phase control that is independent of quorum sensing, unlike other virulence factors of V. cholerae. Additionally, in stationary-phase cultures, all RTX toxin activity is associated with bacterial membranes or outer membrane vesicles.


arrow
INTRODUCTION
 
Oral ingestion of Vibrio cholerae by human hosts via contaminated food and water can result in cholera, a severe diarrheal disease that can quickly cause death if left untreated (34). The secreted ADP-ribosylating cholera toxin (CT) causes the profuse diarrhea that is a hallmark of cholera disease and is one of the main virulence factors produced by the bacterium. V. cholerae also expresses the toxin-coregulated pilus (TCP), which is essential for sustained colonization of the human small intestine. In addition, hemagglutinin/protease (HAP), hemolysin, and RTX toxin have been identified as secreted "accessory" toxins that also contribute to disease pathogenesis (11).

RTX toxin is a very large toxin with a deduced molecular mass of 484 kDa and causes cell rounding and the depolymerization of the actin cytoskeleton in a broad range of cell types. Concurrent with actin stress fiber disassembly, actin monomers are covalently cross-linked into dimers, trimers, and higher multimers which can be visualized by Western blotting (13, 25). rtxA, the 13,635-bp gene encoding the RTX toxin, is the largest open reading frame of the V. cholerae genome (17, 25). rtxA is found in both clinical and environmental isolates of V. cholerae, but not in O1 classical biotypes (7, 10, 25).

V. cholerae RTX toxin is a member of the RTX (repeat-in-toxin) family of toxins (25) that includes Escherichia coli hemolysin (HlyA) and Bordetella pertussis adenylate cyclase-hemolysin toxin. All RTX exoproteins are exported out of the cell by type I secretion systems (T1SS) (41). These secretion systems commonly consist of three components: an inner membrane transport ATPase, a transmembrane linker protein, and an outer membrane porin, exemplified by the E. coli hemolysin T1SS consisting of HlyB, HlyD, and TolC, respectively (1, 19). V. cholerae RTX toxin is secreted by an atypical T1SS that is composed of four components: RtxB, RtxD, TolC, and a second ATPase RtxE (5).

With the exception of tolC, the genes necessary for RTX toxin production and secretion are found in the rtx locus, which is tightly linked to the CTX{Phi} integration site (25). The six genes of the locus are organized in two divergently transcribed operons (Fig. 1A). The rtxA operon consists of the RTX toxin gene rtxA downstream of the putative RTX activating acyltransferase gene rtxC and a conserved hypothetical gene VC1449. The rtxBDE operon encodes three components of the T1SS. The organization depicted in Fig. 1A is conserved among a subset of very large RTX exoproteins (5) but differs from the single-operon organization of the E. coli hly locus.


Figure 1
View larger version (32K):
[in this window]
[in a new window]

  		FIG. 1. The rtx locus of V. cholerae contains six genes in two divergently transcribed operons. (A) Schematic illustration of the rtx locus of V. cholerae. CHP, conserved hypothetical protein. (B) The rtx intergenic region contains 411 bp that lie between the rtx operon and the rtxBDE operon. The transcriptional start site of the rtxBDE transcript was determined by 5' RACE analysis and is underlined. The putative –35 and –10 sites are underlined and were determined by visual inspection of the promoter region. The discriminator region is double underlined. Bases corresponding to promoter trim-backs are in bold and labeled above.

V. cholerae regulates the two main virulence factors TCP and CT as well as HAP via quorum sensing, a system that allows bacteria to communicate and coordinate gene expression in a cell density-dependent manner via extracellular chemical signaling molecules (31, 44). CT and TCP are both expressed at low cell densities and repressed at high cell densities, while HAP and the PrtV protease are repressed at low cell densities and upregulated during stationary phase (43).

Previous studies have identified that RTX activity is detectable only in supernatant fluids from log phase cultures (5, 13). In this work, we have determined that multiple mechanisms coordinately contribute to growth phase regulation. This study focuses on proteases, which inactivate secreted RTX toxin as well as initial studies of growth phase regulation of the atypical T1SS encoded by rtxBDE. The regulation of rtxBDE is found to be at the transcription level by a mechanism that is independent of the three identified quorum-sensing systems of V. cholerae.


arrow
MATERIALS AND METHODS
 
Bacterial strains and growth conditions. The bacterial strains used in this study and relevant genotypes are listed in Tables 1 and 2. V. cholerae was grown at 30°C in Luria broth (LB) supplemented with streptomycin (100 µg/ml) and kanamycin (50 µg/ml) as necessary.


View this table:
[in this window]
[in a new window]

  		TABLE 1. RTX toxin activity and protease activity in supernatant fluids of various V. cholerae strainsa


View this table:
[in this window]
[in a new window]

  		TABLE 2. RTX toxin activity in supernatant fluids and cell lysates of various V. cholerae strains

Generation of mutations in V. cholerae strains. The N16961{Delta}luxO mutant was constructed by the method of Skorupski and Taylor (37). BMV5 (Bah1 {Delta}toxS) was made by sucrose-dependent homologous recombination with the sucrose-counterselectable plasmid pBM8, which contains a 341-bp deletion within toxS created by restriction enzyme digestion of internal SalI and XhoI sites. All {Delta}hapA mutations were constructed with the counterselectable pCVD{Delta}HapSal1 as previously described (12). Counterselectable plasmids were introduced into V. cholerae by conjugation with SM10{lambda}pir on an LB agar plate. Cointegrants were subjected to sucrose-dependent counterselection as previously described (12), and isolated colonies were screened for the presence of a gene deletion by direct colony PCR across the deletion junction and by loss of protease activity on 1% milk agar plates.

Recombinant methods and Western blotting. All recombinant DNA methods were performed as previously described by Sambrook and Russell (35) using reagents from DNA purification kits purchased from QIAGEN (Valencia, CA) when appropriate. PCR was performed using Vent DNA polymerase or Pfx DNA polymerase according to manufacturers' protocols. Enzymes were obtained from Invitrogen (Carlsbad, CA) or New England Biolabs (Beverly, MA). PCR primers were obtained from Integrated DNA Technologies (Coralville, IA) or Sigma Genosys (The Woodlands, TX), and the sequences are available upon request. Automated DNA sequencing was performed at either the Northwestern University Biotechnology Laboratory or the University of Chicago Cancer Research Center DNA Sequencing Facility. For Western blotting, sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) gels were transferred to nitrocellulose in a Bio-Rad Mini Trans-Blot transfer cell at room temperature. Protein bands were detected using the appropriate primary antibody, a 1:5,000 anti-rabbit immunoglobulin G-peroxidase conjugate (Jackson ImmunoResearch Laboratories, West Grove, PA) secondary antibody, and ECL reagents.

Preparation of supernatant fluids. V. cholerae cultures were grown at 30°C to the desired optical density. Bacteria were pelleted by centrifugation, and supernatant fluid was filtered over a 0.2-µm Corning Spin-X microfilter.

Preparation of bacterial cell lysates and separation of cellular fractions. Bacteria from 25-ml cultures grown to log phase (A600 = 0.4) or an equivalent number of bacteria from stationary phase were collected by centrifugation at 15,000 x g for 5 min. Pellets were washed in 20 ml 2 mM CaCl2. Pellets were resuspended in 6.4 ml ice-cold distilled H2O and sonicated with a Bronson digital Sonifier 450 with a 3-mm tapered microtip at 30% amplitude and 50% duty for 40 s. After sonication, the lysate was adjusted to 20 mM Tris, 100 mM NaCl, 1 mM EDTA, pH 8.0, by the addition of 10x stock buffer. The lysate was centrifuged first at 5,000 x g to pellet unlysed bacteria and then at 20,000 x g to pellet debris. The supernatant was retained as total cell lysate. To separate membranes from cytoplasm, total cell lysate was centrifuged at 100,000 x g for 30 min. The supernatant was retained as the cytoplasmic fraction, and the pellet, representing the membrane fraction, was resuspended in an equal volume of 20 mM Tris, 100 mM NaCl, 1 mM EDTA, pH 8.0. The separation of whole-cell lysates into cytoplasmic and membrane fractions was verified by ß-galactosidase assays (29) and Western blots for the inner membrane protein EpsL using a rabbit polyclonal EspL antibody (M. Sandkvist, University of Michigan).

Actin cross-linking assay. A total of 0.5 ml of the sample to be tested was added to 4.5 ml Dulbecco's modified eagle medium without serum and added to 2.5 x 105 HEp-2 cells. Unless indicated otherwise, HEp-2 cells were collected after 90 min for the detection of actin cross-linking by Western blotting as previously detailed (36).

Zymogram. To prepare concentrated supernatant fluids, 20-ml cultures were grown overnight and centrifuged at 4°C. Supernatant fluids were placed in prewet 1-kDa-cutoff dialysis tubing and then set in polyethylene glycol 3000 at 4°C. Over several hours, fresh polyethylene glycol 3000 was added and supernatant fluids were allowed to dehydrate until 2 to 3 ml of each sample remained. Protein concentrations were determined by bicinchoninic acid protein assay (Pierce, Rockford, IL), and protease activity was measured by a modified skim milk contact zymogram (18). Briefly, 5 µg of concentrated supernatant fluids was run on a 10% SDS-PAGE gel. SDS was removed by washing the gel with 2.5% Triton X-100 at room temperature for 30 min, and the gel was then equilibrated with 0.1 M glycine-NaOH, pH 8.3, for 5 min at room temperature. The gel was then overlaid with a 10% skim milk gel for 1 h at 37°C. The overlay gel was then stained with Coomassie brilliant blue 250 for 1 h, and protease activity was viewed as clear bands on the blue background.

Protease activity measured by azocasein assay. Protease activity was measured using a modified azocasein hydrolysis assay (38). Briefly, 50 µl of filtered supernatant fluid was added to 800 µl of azocasein at 2 mg/ml in 0.1 M Tris-HCl, pH 8.0. After incubation at 37°C, the reaction was stopped by adding 160 µl of 50% trichloroacetic acid to each tube. Samples were immediately centrifuged at 16,000 x g for 15 min at room temperature. The absorbance of the solution was read at 366 nm on a Beckman DU Series 500 spectrophotometer (Fullerton, CA).

Generation of anti-RtxE antibody. The RtxE peptide TRFTTDAHRLAEQTLPQEC was synthesized and used to produce an anti-RtxE antibody in rabbits by standard protocols (Proteintech Group, Inc., Chicago, IL).

Construction of lacZ reporter plasmids. The full-length rtxBDE promoter::lacZ reporter plasmid pBB24 was created by using primers rtx pro 2 and rtxB HindIII to amplify the rtxBDE promoter region of the V. cholerae strain N16961 and add a HindIII restriction site to aid in subcloning. The promoter region was amplified from 386 bp upstream of the rtxB ATG start site to 6 bp downstream of the ATG start site. The PCR product was cloned into pCR BluntII-TOPO (Invitrogen) and sequenced. lacZ was fused in frame to the rtxBDE promoter as a HindIII fragment. Subsequent trim-back deletions were constructed by using pBB24 as a PCR template for primer lacZ, which binds 453 bp into the lacZ gene and various primers that bound successively closer to the ATG start site of rtxB. PCR products were cloned into pCR BluntII-TOPO, the partial fragment of lacZ was removed with a HindIII digestion, and full-length lacZ was inserted as a HindIII fragment. The Stratagene QuikChange kit (La Jolla, CA) was used for site-directed mutagenesis of the discriminator region downstream of the rtxBDE –10 sequence to introduce nucleotide changes from GCCG to CGAT and to create a ClaI site used to screen for the presence of the mutation. All promoter fragments and junctions were sequenced to assure that mutations were not introduced by PCR.

ß-Galactosidase assays. V. cholerae strains containing lacZ reporter plasmids were grown at 30°C in LB. Samples were withdrawn at indicated optical densities and assayed for ß-galactosidase activity by the method of Miller (29).

RNA isolation and generation of cDNA. Overnight cultures of wild-type strain N16961 were diluted 1:100 in fresh LB and grown at 30°C. RNA was isolated using the QIAGEN RNeasy purification kit and treated with DNase (Promega, Madison, WI), as per the manufacturer's instructions. cDNA was synthesized as follows: 5 µg total RNA and 30 µg random hexamer primers (Roche, Indianapolis, IN) were combined and denatured at 70°C for 10 min and then placed on ice for 10 min. Dithiothreitol, dNTP mixture, RNasin (Promega), SuperScriptII (Invitrogen), and H20 were added to a final volume of 50 µl. Reverse transcription was carried out at 42°C for 2.5 h, and the SuperScript was inactivated by heating at 94°C for 10 min. The cDNA was treated with RNaseI (Promega) and was then further purified and concentrated using the QIAGEN PCR purification kit. A reverse transcriptase-free reaction was included as a control to exclude chromosomal DNA contamination.

Reverse transcriptase PCR (RT-PCR). Seventy-seven nanograms of cDNA, indicated primers, and TaqPro Red Complete DNA polymerase (Denville Scientific, South Plainfield, NJ) were mixed in 25-µl reactions. cDNA was amplified during 30 cycles of PCR in a PTC-200 Peltier thermal cycler. Transcripts were then analyzed by agarose gel electrophoresis.

Quantitative PCR. Quantitative PCR (Q-PCR) was performed in 20-µl reactions using 39 ng of each cDNA sample, Brilliant SYBR green QPCR Master Mix (Stratagene), and the specific primer pair. Template-free or reverse transcriptase-free reactions were included as controls. All PCRs were performed with the MJ Research Opticon Chromo 4 (Bio-Rad, Richmond, CA). The PCRs for each independent experiment were run in duplicate to check the consistency of the PCRs. recA was used as an internal control for all of the PCRs, and transcript amounts for all of the strains and time points were calculated by the following equation: relative gene expression Formula, where CT is the threshold cycle.

RACE analysis. RNA was isolated from strain N16961 grown in LB at 30°C to log phase (A600 = 0.4) and reverse transcribed using the primer rtxB8. cDNA was purified and tailed with poly(dC) using a 5' rapid amplification of cDNA ends (RACE) system kit (Invitrogen). The rtxB 5' end was amplified using the RACE kit AAP primer and rtxB9 primer. The PCR product was run on a 1% agarose gel, a 500-bp band was excised, and the agarose plug was used for a second round of PCR using the RACE kit UAP primer and rtxB9 primer. The reamplified 500-bp product was excised and purified from a 1% agarose gel and sequenced. 5' RACE was performed twice from two RNA preparations to confirm the accuracy of the transcriptional start site.


arrow
RESULTS
 
Proteases present in culture supernatant fluids inactivate RTX toxin. The original report on the discovery of the RTX toxin determined that RTX toxin was not secreted to supernatant fluids (25). These findings were subsequently amended when both RTX toxin and its associated actin cross-linking activity were found in culture supernatant fluids (13), and secretion of the toxin was shown to be dependent upon an atypical T1SS (5). For both studies, the activity in supernatant fluids was limited to Inaba strain N16961 and was not routinely detected in the supernatant fluids of strain Bah1, a derivative of the Ogawa strain E7946 in which there is a deletion of the CTX{Phi} element except the repeat sequence element, thus creating a {Delta}core strain (5, 13; our unpublished observations). The RTX toxin-associated actin cross-linking activity produced by strain KFV43 (N16961{Delta}hapA) was found to be restricted to log phase growth, with maximal activity between 0.4 and 1 at A600 (Fig. 2).


Figure 2
View larger version (22K):
[in this window]
[in a new window]

  		FIG. 2. RTX toxin-associated activities are absent in supernatant fluids from stationary-phase cultures. Supernatant fluids collected from a culture of strain KFV43 (N16961{Delta}hapA) grown to various optical densities were assayed for RTX activity via actin cross-linking assay. LB was used as a mock treatment. Monomer (M), dimer (Di), trimer (Tri), and tetramer (Te) forms of actin are labeled at right. Numbers indicate molecular masses in kilodaltons.

A major difference between N16961 and E7946 is the amount of protease activity found in supernatant fluids due to a mutation in hapR in N16961 (20, 44). The presence of functional or mutated hapR genes has been shown to differ in geographically diverse strains of V. cholerae (20). HapR activates hapA expression, and as shown in Table 1, wild-type E7946 produced 21-fold-greater protease activity at stationary phase compared to wild-type N16961. Consistent with growth phase control of protease production by quorum sensing, the protease activity of E7946 when sampled from stationary-phase cultures was 22-fold higher than that from log phase cultures. RTX toxin activity was absent in supernatant fluids of log phase cultures of E7946, and only nondetectable levels of protease activity correlated with the presence of RTX toxin-associated actin cross-linking activity in supernatant fluids from log phase cultures (Table 1). The deletion of the major protease gene hapA from either N16961 or the E7946 derivative Bah1 to create strain KFV43 or Bah1P, respectively, reduced protease activity in supernatant fluids, and RTX activity was now detectable in log phase cultures of Bah1P (Table 1). However, both {Delta}hapA strains produced measurable protease activity from other less abundant proteases in stationary phase and RTX activity was still not detected (Table 1 and Fig. 3).


Figure 3
View larger version (88K):
[in this window]
[in a new window]

  		FIG. 3. Deletion of toxS alone or in combination with hapA reduces protease activity in strain Bah1. Secreted proteases in overnight culture supernatant fluids from strains Bah1 (E7946{Delta}core), BMV5 (Bah1{Delta}toxS), BMV6 (BMV5{Delta}hapA), and Bah1P (Bah1{Delta}hapA) were detected by skim milk overlay zymogram. White areas on the gel denote protease activity. Numbers indicate molecular masses in kilodaltons.

Additional protease production mutants in both N16961 and E7946 backgrounds were tested to ascertain whether proteases other than HAP account for the absence of RTX activity in stationary phase. Strain E7946mpc, which was generated by Tn5 mutagenesis, has a multiple protease control (mpc) defect (28, 40) and reduced protease activity both in log and stationary-phase growth (Table 1). E7946mpc has transposon insertions in toxS and dcuA, although these mutations are not linked to the protease control defect, which is due to a spontaneous mutation in the regulatory gene luxO (40). Regardless, the deletion of toxS in the Bah1 also created a mutant (BMV5) with the mpc phenotype, indicating that spontaneous mpc defects may commonly arise in toxS mutants. A deletion of toxS in Bah1, either alone or in combination with a deletion in hapA, reduced protease activity as measured both by zymogram analysis (Fig. 3) and azocasein assays (Table 1). For these E7946-derived protease mutants, RTX activity was detected in log phase but remained absent in stationary phase (Table 1).

Similarly, a deletion of the N16961 gene pilD, the pilin processing protein that affects the secretion of proteases by the eps secretion system (12, 26), either alone or in combination with {Delta}hapA, reduced protease activity to nondetectable levels in both log and stationary phase. Yet, RTX activity remained absent in stationary-phase cultures. Thus, measurable protease activity correlates with the inactivation of RTX activity in only log phase, not in stationary phase.

In order to determine whether a level of protease activity undetectable by azocasein assays in KFV18R and KFV44 stationary-phase supernatant fluids could still inactivate RTX activity, these supernatant fluids were coincubated with RTX-containing supernatant fluids prior to the addition to HEp-2 cells to directly assay the toxin-inactivating activity of the supernatant fluids. As expected, the incubation of the RTX-positive fluids with supernatant fluids from protease-producing control strains E7946 and N16961 was sufficient to destroy RTX activity (Fig. 4, lanes 3 and 4), while LB alone had no effect (lane 2). However, coincubation with KFV18R and KFV44 stationary-phase supernatant fluids did not inactivate the RTX activity, which was consistent with the lack of protease activity in these cultures (lanes 5 and 6). These data demonstrate that the absence of RTX toxin activity in supernatant fluids during stationary phase is not always correlated with protease expression, indicating that RTX inactivation by proteases is not solely responsible for the lack of RTX activity in stationary-phase cultures. Thus, another mechanism must also be functioning to account for the lack of RTX toxin activity in supernatant fluids of strains that do not produce extracellular proteases. For the remaining studies, we used KFV43 as this strain had the lowest protease activity in supernatant fluids (Table 1) but does not have the uncharacterized growth defect suppressor mutation of KFV18R or KFV44 (12).


Figure 4
View larger version (27K):
[in this window]
[in a new window]

  		FIG. 4. Protease-containing supernatant fluids inactivate RTX toxin activity. Supernatant fluids were collected and filtered to remove bacteria. A total of 0.25 ml of filtered supernatant fluid containing active RTX toxin (from log phase [A600 = 0.4] E7946mpc cultures) was mixed with 0.25 ml of LB or supernatant fluid from overnight cultures of the indicated strain and incubated at 37°C for 30 min. Treated supernatant fluids were added to HEp-2 cells, and cells were processed for the detection of actin cross-linking. LB was used as a mock treatment. Monomer (M), dimer (Di), trimer (Tri), and tetramer (Te) forms of actin are labeled at right. Numbers indicate molecular masses in kilodaltons.

Production of the RTX toxin T1SS is regulated by growth phase. In a recent study, we demonstrated that during exponential growth phase, mutants in the T1SS for the export of RTX toxin accumulate active toxin intracellularly (5). Therefore, if toxin is expressed in stationary phase but not secreted, active toxin should be present in cell lysates. As shown in Fig. 5, only supernatant fluids from log phase cultures contain RTX toxin activity, but clarified cell lysates contain active toxin in both log and stationary phases of growth. Initially, these data suggested that the secretion of RTX toxin, not toxin expression, is repressed during stationary phase. Other V. cholerae {Delta}hapA isolates showed a similar pattern of RTX toxin activity, demonstrating that the growth phase control of the T1SS is broadly applicable (Table 2).


Figure 5
View larger version (19K):
[in this window]
[in a new window]

  		FIG. 5. RTX toxin-associated activities are absent in supernatant fluids but present in cell lysates from stationary-phase cultures. Equivalent numbers of bacteria of strain KFV43 (N16961{Delta}hapA) grown to various cell densities were collected and lysed. Filtered supernatant fluids or cell lysates were then evaluated for RTX activity by actin cross-linking assay. Luria broth and 2 mM Tris, 1 mM EDTA, pH 8.0 (TE), were used as mock treatments. Monomer (M), dimer (Di), trimer (Tri), and tetramer (Te) forms of actin are labeled at right. Numbers indicate molecular masses in kilodaltons.

The T1SS for RTX toxin is composed of four proteins, RtxB, RtxD, RtxE, and TolC (5). A peptide antibody was raised against the ATPase RtxE, which has a predicted molecular mass of 79 kDa. This antibody was then used to perform Western blotting to detect RtxE in bacteria. The RtxE antibody is specific for the ATPase RtxE and not the ATPase RtxB, as shown by the absence of the RtxE band in a strain containing an in-frame deletion of rtxE (CW149) and in strain BBV16, which has a polar insertion of a kanamycin resistance cassette in the upstream gene rtxB inactivating the entire rtxBDE operon (Fig. 6A). The RtxE band is detectable in stain BBV21, which contains a nonpolar deletion of rtxB. RtxE is detected in strain KFV43 grown to log phase, but its expression is significantly reduced during stationary phase. Thus, the expression of the secretion apparatus for RTX toxin is under growth phase control and is repressed in stationary phase.


Figure 6
View larger version (29K):
[in this window]
[in a new window]

  		FIG. 6. The T1SS for export of V. cholerae RTX toxin is regulated by growth phase. (A) Equivalent numbers of bacteria from overnight cultures or log phase cultures (A600 = 0.4) of V. cholerae El Tor strain N16961 (wild type), BBV16 (N16961 {Delta}hapA rtxB::Km), BBV21 ({Delta}hapA {Delta}rtxB), or CW149 ({Delta}hapA {Delta}rtxE) were collected and boiled in 1x SDS buffer. Proteins were separated on an 8% SDS-PAGE gel, and RtxE was subsequently detected by Western blotting. (B) Agarose gel analysis of RT-PCR of the rtxB, rtxD, and rtxE genes that encode the T1SS. RNA was isolated from wild-type N16961 grown to the indicated optical densities. recA was used as an internal control. (C) Relative expression of T1SS gene mRNA transcripts in log (A600 = 0.4) and stationary (A600 = 5) phases. Values are representative of at least two experiments.

Growth-phase control of rtxBDE expression is at the level of gene transcription. In order to ascertain whether the T1SS is regulated at the transcriptional level, RT-PCR was used to detect T1SS gene transcripts in RNA isolated from a wild-type N16961 culture at various time points during the growth cycle. rtxB, rtxD, and rtxE transcripts were detected 30 min (A600 = 0.1) after the N16961 overnight culture was subcultured in fresh LB, but the T1SS gene transcripts were barely detectable after 6 h of growth (A600 = 5) relative to the internal control recA (Fig. 6B). Q-PCR showed that the relative expression levels of T1SS genes in log phase (A600 = 0.4) were at least 10 times higher than the relative expression levels in stationary phase (A600 = 5) (Fig. 6C). rtxB transcript levels were consistently higher than rtxD and rtxE levels in log phase cultures, which is not surprising since rtxB would be the first gene in the operon to be transcribed. These data show that the T1SS is regulated by growth phase and the level of control most likely occurs at the level of gene transcription or mRNA stability.

To demonstrate that the regulation of the T1SS is through promoter regulation, the rtxBDE promoter was fused to the V. cholerae lacZ gene at the ATG site of rtxB in the multicopy plasmid pCR BluntII-TOPO. ß-galactosidase activity from the plasmid was monitored over the growth cycle in N16961{Delta}lacZ (Fig. 7). Monitoring of growth (Fig. 7A) showed that N16961{Delta}lacZ carrying the reporter plasmid grew exponentially for 4 h with a doubling time of 24 min. At an A600 of 1.3, cell division slowed to a doubling time of 56 min for 5 h until the culture reached an average A600 of 6 when cell growth ceased. The expression of lacZ in this culture was growth phase regulated with maximal expression of ß-galactosidase at an A600 of 0.4 (Fig. 7B) and dropped by over 50% when cell growth slowed at an A600 congruent to 1. ß-galactosidase activity was not present with a control plasmid containing lacZ, but none of the rtxBDE promoter sequence. These data showed that the genes encoding the T1SS are downregulated during stationary phase, and regulation is controlled by the rtxBDE promoter.


Figure 7
View larger version (17K):
[in this window]
[in a new window]

  		FIG. 7. ß-galactoside activity produced by an N16961 strain containing an rtxBDE promoter::lacZ reporter plasmid peaks during log growth. (A) Growth curves of strains assayed for ß-galactosidase activity. (B) N16961{Delta}lacZ strains containing a full-length rtxBDE promoter::lacZ reporter plasmid or a negative control plasmid containing only lacZ and no portions of the rtxBDE promoter (no pro) were assayed for ß-galactosidase activity at various time points.

Identification of the rtxBDE transcriptional start site. To further understand the growth phase control of the rtxBDE promoter, the start site for the transcription of the operon was mapped using the Invitrogen 5' RACE kit. The +1 start site was mapped to a thymine residue 37 bp upstream of the rtxB ATG (Fig. 1B). The proposed –10 sequence TAGGAT shows two mismatches from the consensus TATAAT, and the putative –35 sequence TTGAAC shows two mismatches from the consensus TTGACA (8). An interesting feature of the rtxBDE promoter is the presence of an extremely G+C-rich 7-bp region between the –10 sequence and the transcription start site, which is often called the discriminator region (Fig. 1B).

Functional characterization of the rtxBDE promoter. In order to ascertain how much of the promoter region is essential for growth phase control, trim-back deletions of the rtxBDE promoter::lacZ reporter were constructed and assayed for ß-galactosidase activity (Table 3). A deletion of 300 bp, leaving only 13 bp upstream of the putative –35 site still exhibited growth phase control. ß-galactosidase activity was absent either in a no-insert control or when the rtxBDE promoter was deleted to 36 or 17 bp upstream of the transcriptional start site (Fig. 7B; Table 3).


View this table:
[in this window]
[in a new window]

  		TABLE 3. ß-galactosidase activity of trim-back deletions of the rtxBDE promoter::lacZ reporter plasmid

Transcriptional regulation by activation generally requires promoter elements upstream of the –35 site as binding sites for transcriptional activators. Although only 13 bp upstream of the putative rtxBDE –35 site were required for ß-galactosidase activity from the rtxBDE promoter::lacZ reporter plasmid, it is possible that a positive regulatory element binds this region during log phase growth. However, the minimal requirements for the growth phase-regulated rtxBDE promoter suggests it is likely that the promoter is regulated by a repressive element activated during late log phase.

Growth-phase control of rtxBDE expression is not linked to quorum sensing. A common growth phase-dependent repression mechanism is quorum sensing. In V. cholerae, both CT and TCP production are repressed in stationary phase via quorum-sensing-dependent regulation (30, 44). The master regulator of quorum sensing in V. cholerae is the transcriptional regulator HapR, which represses the expression of genes required for CT and TCP gene activation during stationary phase (44). Whole-genome sequence analysis (17) revealed that N16961 has a defective quorum-sensing system due to a naturally occurring frameshift mutation in hapR that introduces an opal codon (44). DNA sequencing of this region amplified by PCR verified that our laboratory isolate contains this mutation, indicating that if quorum sensing is regulating rtxBDE expression, it occurs at a signaling branch upstream of HapR.

The three known quorum-sensing pathways of V. cholerae converge at the response regulator LuxO, which operates upstream of HapR in the quorum-sensing pathway (22, 31, 43). A deletion in luxO had no effect on the growth phase control of the rtxBDE::lacZ reporter (Table 3), consistent with microarray studies which did not report significant changes in rtxB, rtxD, or rtxE expression in luxO mutants in strains with active quorum-sensing systems (44). Since neither hapR nor luxO is required for growth phase regulation, it is unlikely that rtxBDE is linked to any of the three identified quorum-sensing pathways, although it could be linked to an unidentified response regulator other than luxO.

Growth-phase regulation is determined by the promoter discriminator region. The V. cholerae genome is only 47% G+C, therefore we wanted to determine whether the heptanucleotide G+C-rich discriminator region (GCCGCCC) downstream of the –10 sequence is required for growth phase control. The GCCG bases immediately downstream of the –10 sequence were mutated to CGAT in the full-length rtxBDE promoter::lacZ reporter plasmid. During log phase growth, the ß-galactosidase activity of the discriminator mutant reporter plasmid was similar to that of the full-length rtxBDE promoter::lacZ reporter plasmid (Table 3). However, in stationary phase, the discriminator mutant reporter plasmid did not have a reduction in ß-galactosidase activity, which demonstrates that these bases are critical for growth phase control but not for promoter function. Additionally, the alteration to constitutive ß-galactosidase expression from the discriminator mutant reporter plasmid demonstrates that the discriminator region of the rtxBDE promoter is required for negative regulation during stationary phase.

The rtxA toxin gene is also regulated by growth phase. Based on data presented thus far, a model was proposed in which RTX toxin is expressed constitutively, but is secreted by the T1SS only during log growth (4). However, it seemed unlikely that RTX toxin would continue to accumulate in the bacterial cytoplasm during stationary phase due to the presumably extensive energy required to translate this extremely large toxin. Therefore, RT-PCR was used to detect rtxA transcripts in RNA isolated from a wild-type N16961 culture at various time points during the growth cycle. Counter to the accumulation model supported by Fig. 5, PCR directed against rtxA cDNA demonstrated that rtxA transcripts are greatly reduced during stationary phase (Fig. 8A). In stationary-phase (A600 = 5), rtxA transcripts are decreased 10-fold compared to that in log phase (A600 = 0.4) as determined by Q-PCR (data not shown). Thus, the RTX toxin gene itself is apparently regulated by growth.


Figure 8
View larger version (42K):
[in this window]
[in a new window]

  		FIG. 8. RTX toxin is regulated by growth phase, and activity in stationary-phase cultures is associated with the bacterial membrane. (A) Agarose gel analysis of RT-PCR of rtxA transcripts in RNA isolated from wild-type N16961 grown to the indicated optical densities. recA was used as an internal control. (B) HEp-2 cells were treated with filtered supernatant fluids (sup), total cell lysates (total lys), cytoplasm (cyto), or membrane (mem) fractions from log (A600 = 0.4) or stationary-phase cultures (A600 = 3.0) of strain KFV43 (N16961 {Delta}hapA) or BBV16 (N16961 {Delta}hapA, rtxB::Km). LB was used as a mock treatment. Monomer (M), dimer (Di), trimer (Tri), and tetramer (Te) forms of actin are labeled at right. (C) The separation of total cell lysate into cytoplasmic and membrane fractions was ensured by assaying for the cytoplasmic marker ß-galactosidase, shown in Miller units (MU), and by Western blotting for the inner membrane protein EspL. Numbers indicate molecular masses in kilodaltons.

The RNA-binding protein Hfq has been shown to be involved in the regulation of V. cholerae virulence factors (23), and it has been reported that an hfq mutant has increased expression of rtxA and rtxC in log phase (9). RNA transcripts from this hfq mutant strain were analyzed by Q-PCR, and VC1449, rtxC, and rtxA were found to be downregulated in the hfq mutant during stationary phase to a level similar to that of the wild-type strain (data not shown). Therefore, growth phase control of RTX toxin is apparently not linked to Hfq, and the mechanism of gene suppression remains unknown.

RTX toxin activity is associated with the membranes of stationary-phase bacteria. To account for the discrepancy wherein rtxA transcripts are greatly reduced in stationary phase (Fig. 8) but RTX toxin activity is not decreased in cell lysates (Fig. 5), two possible models were considered. In the first model, the toxin would be extremely stable when held in the cytoplasm and, thus, active protein would remain present in the stationary-phase bacteria. In the second model, secreted toxin would attach to the outer membrane or outer membrane vesicles and remain protected from proteolysis and thus retain activity through stationary phase. To distinguish these models, cell lysates from stationary-phase cultures of the T1SS mutant BBV16 were tested for the presence of RTX toxin activity. No toxin activity was present in cell lysates, indicating that the toxin is not unusually stable when accumulating in the cytoplasm and is likely rapidly turned over in the absence of a secretion apparatus (Fig. 8B).

To determine whether the active toxin detected in cell lysates from stationary-phase cultures of strain KFV43 (N16961{Delta}hapA) is associated with bacterial membranes or outer membrane vesicles, whole bacteria grown to stationary phase were sonicated to obtain total cell lysates, which were subsequently centrifuged at 100,000 x g to pellet the membrane fraction. While RTX toxin activity was detected in total cell lysates, toxin activity was associated with only the membrane fraction, not the cytoplasmic fraction (Fig. 8B). Due to the fact that RTX activity was detected in only the cytoplasmic fraction of log phase BBV16 cultures and not in the membrane fraction, the large toxin was not simply pelleted during the high-speed centrifugation. Thus, in stationary phase, RTX toxin is not produced, but activity remains in cell lysates due to its association with the bacterial membrane. Increased attachment of bacteria with HEp-2 cells by low-speed centrifugation did not restore actin cross-linking activity to stationary-phase cultures, indicating that bacterial membrane associated toxin is not biologically active unless sheared from the bacterium by sonication (data not shown).


arrow
DISCUSSION
 
In this study, we have presented data that describes two control mechanisms for the regulation of RTX activity: proteolytic degradation of RTX toxin and the regulation of the T1SS that exports the toxin. In addition, we find that the RTX toxin itself is growth phase regulated by an as-yet-uncharacterized mechanism. Based on these results, we put forth a three-stage model to explain the growth phase regulation of V. cholerae RTX toxin.

During exponential growth, the rtxA and rtxBDE operons are expressed and the atypical T1SS encoded by rtxBDE and tolC exports RTX toxin into the extracellular milieu, where the toxin can subsequently act on host cells (Fig. 9A, B, and C). When bacteria begin to enter stationary phase, the RtxBDE T1SS is no longer produced due to transcriptional regulation via what is most likely a repressive mechanism (Fig. 9A). Consequently, in the absence of the secretion apparatus, any RTX toxin remaining in the bacterium would not be secreted and is likely degraded. rtxA expression is likewise downregulated in stationary phase via an unknown mechanism, and thus, no additional RTX toxin would be produced. Concurrent with RTX toxin and T1SS repression, the expression of proteases is upregulated during stationary phase, which destroys the activity of previously secreted RTX toxin (Fig. 9E), although toxin associated with the membrane or outer membrane vesicles could be protected (Fig. 9D). The increase in protease expression occurs in both quorum-sensing-positive and -negative strains, although the quorum-sensing-positive strain E7946 has a more robust increase in protease expression due to massive upregulation of HAP in stationary phase. Consequently, in certain strains of V. cholerae, quorum sensing indirectly regulates RTX activity.


Figure 9
View larger version (27K):
[in this window]
[in a new window]

  		FIG. 9. Model of growth phase regulation of RTX toxin export as described in the text.

A major question raised by this study is what is the nature of the putative repressor that regulates rtxBDE? We hypothesize that this repressor lies outside of the rtx locus due to the fact that the rtxBDE::lacZ reporter showed growth phase control when the plasmid was introduced into the N16961-derived strain KFV80 or the classical strain O395, which both contain a large deletion of the rtx locus ({Delta}rtxACBD) (15, 27; data not shown). We have also shown that this repressor is not directly linked to quorum sensing.

Indeed, it is more reasonable to consider that V. cholerae may couple the regulation of the rtx locus to the detection of stress since the locus represents 1% of the large chromosome and would require extensive energy to produce, a high cost during periods of stress. Altered expression of multiple rtx locus transcripts in microarray studies of rpoN and rpoS mutants also indicate a linkage to stress responses (42). In our study, the G+C-rich discriminator region of the rtxBDE promoter was shown to be essential for growth phase control. Studies in E. coli have demonstrated that G+C-rich discriminator regions are hallmarks of promoters that are negatively controlled by the stringent response alarmone (p)ppGpp (6, 16, 39) and that (p)ppGpp concentrations increase with the onset of stationary phase (32). Preliminary studies on the structure of the rtxA promoter also indicate that a G+C-rich discriminator is present in the promoter that regulates VC1449, rtxC, and rtxA (our unpublished observations). Therefore, the T1SS genes as well rtxA could potentially be negatively regulated during stationary phase via (p)ppGpp. Further studies will be required to determine the exact nature of the growth phase-dependent mechanism(s) by which the T1SS and RTX toxin are regulated.

Another interesting finding of this work is that all RTX activity in stationary-phase cultures was found to be associated with the bacterial membrane fraction, although its subcellular localization remains unknown. Proteins secreted via T1SS are exported without the formation of periplasmic intermediates; thus, RTX toxin would not likely be located in the periplasmic space of the bacterium (21). It has been shown in E. coli that HlyA requires the ATPase HlyB in order to associate with the inner membrane prior to export, suggesting that cytoplasmic-localized RTX toxin is peripherally associated with the RTX T1SS ATPases, accounting for the membrane localization in log phase cultures (14). However, we have shown that the ATPase RtxE is significantly reduced in stationary-phase cultures of V. cholerae and the second ATPase, RtxB, is likely turned over as well; thus, it is unlikely that RTX toxin associates with the inner membrane by association with T1SS ATPases during stationary phase. Therefore, we hypothesize that RTX toxin is exported during exponential growth and then associates with the bacterial outer membrane, possibly coupled with the outer membrane porin TolC and/or inserted into the outer membrane. RTX toxin could also be released in outer membrane vesicles. Several RTX toxins, including E. coli HlyA and Bordetella pertussis adenylate cyclase toxin, have been found to be associated with the bacterial outer membrane (15, 33). Similar to B. pertussis, spinning stationary-phase V. cholerae bacteria onto HEp-2 cells did not lead to cell rounding or actin cross-linking, thus indicating that membrane-bound RTX toxin is not active unless released from the membrane by sonication. It was recently discovered that E. coli HlyA is also released in outer membrane vesicles (2). Future studies may elucidate whether this phenomenon occurs in V. cholerae with RTX toxin as well.


arrow
ACKNOWLEDGMENTS
 
We thank M. Sandkvist for generously supplying the EpsL antibody. We thank J. Mekalanos and M. Waldor for strains and K. Skorupski for the pKAS32{Delta}luxO construct. We thank B. Billips and M. Rohrer for assistance with RT-PCR.

This work was supported by Public Health Services Grant AI051490 (to K.J.F.S.), a Biomedical Research Support Grant from the Howard Hughes Medical Institute (to K.J.F.S.), and a Pathogenesis of Infectious Disease Award from the Burroughs Wellcome Fund (to K.J.F.S.).


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: Department of Microbiology-Immunology, Northwestern University Feinberg School of Medicine, Tarry 3-713, 303 E. Chicago Avenue, Chicago, IL 60611. Phone: (312) 503-2162. Fax: (312) 503-1339. E-mail: k-satchell{at}northwestern.edu. Back

{triangledown} Published ahead of print on 22 December 2006. Back

{dagger} Present address: Department of Molecular Biology and Microbiology, Tufts University School of Medicine, Boston, MA 02111. Back


arrow
REFERENCES
 
    1
  1. Andersen, C., C. Hughes, and V. Koronakis. 2000. Chunnel vision. Export and efflux through bacterial channel-tunnels. EMBO Rep. 1:313-318.[CrossRef][Medline]
    2. 2
  2. Balsalobre, C., J. M. Silván, S. Berglund, Y. Mizunoe, B. E. Uhlin, and S. N. Wai. 2006. Release of the type I secreted {alpha}-haemolysin via outer membrane vesicles from Escherichia coli. Mol. Microbiol. 59:99-112.[CrossRef][Medline]
    3. 3
  3. Benítez, J. A., L. García, A. Silva, H. García, R. Fando, B. Cedré, A. Pérez, J. Campos, B. L. Rodríguez, J. L. Pérez, T. Valmaseda, O. Pérez, M. Ramírez, T. Ledón, M. D. Jidy, M. Lastre, L. Bravo, and G. Sierra. 1999. Preliminary assessment of the safety and immunogenicity of a new CTX{phi}-negative, hemagglutinin/protease-defective El Tor strain as a cholera vaccine candidate. Infect. Immun. 67:539-545.[Abstract/Free Full Text]
    4. 4
  4. Boardman, B. K., and K. J. Satchell. 2005. The Atypical TISS for secretion of Vibrio cholerae RTX toxin is regulated by growth phase, abstr. B-280, p. 81. Abstr. 105th Gen. Meet. Am. Soc. Microbiol., Atlanta, GA.
    5. 5
  5. Boardman, B. K., and K. J. Satchell. 2004. Vibrio cholerae strains with mutations in an atypical type I secretion system accumulate RTX toxin intracellularly. J. Bacteriol. 186:8137-8143.[Abstract/Free Full Text]
    6. 6
  6. Cashel, M., D. R. Gentry, V. J. Hernandez, and D. Vinella. 1996. The stringent response, p. 1458-1496. In F. C. Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology, 2nd ed., vol. 1. ASM Press, Washington, DC.
    7. 7
  7. Chow, K. H., T. K. Ng, K. Y. Yuen, and W. C. Yam. 2001. Detection of RTX toxin gene in Vibrio cholerae by PCR. J. Clin. Microbiol. 39:2594-2597.[Abstract/Free Full Text]
    8. 8
  8. Choy, H., and S. Adhya. 1996. Negative control, p. 1287-1299. In F. C Neidhardt, R. Curtiss III, J. L. Ingraham, E. C. C. Lin, K. B. Low, B. Magasanik, W. S. Reznikoff, M. Riley, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and Salmonella: cellular and molecular biology, 2nd ed, vol. 1. ASM Press, Washington, DC.
    9. 9
  9. Ding, Y., B. M. Davis, and M. K. Waldor. 2004. Hfq is essential for Vibrio cholerae virulence and downregulates sigma expression. Mol. Microbiol. 53:34.
    10. 10
  10. Faruque, S. M., N. Chowdhury, M. Kamruzzaman, M. Dziejman, M. H. Rahman, D. A. Sack, G. B. Nair, and J. J. Mekalanos. 2004. Genetic diversity and virulence potential of environmental Vibrio cholerae population in a cholera-endemic area. Proc. Natl. Acad. Sci. USA 101:2123-2128.[Abstract/Free Full Text]
    11. 11
  11. Fullner, K. J. 2003. Toxins of Vibrio cholerae: consensus and controversy, p. 481-502. In G. Hecht (ed.), Microbial pathogenesis and the intestinal epithelial cell. ASM Press, Washington, DC.
    12. 12
  12. Fullner, K. J., and J. J. Mekalanos. 1999. Genetic characterization of a new type IV-A pilus gene cluster found in both classical and El Tor biotypes of Vibrio cholerae. Infect. Immun. 67:1393-1404.[Abstract/Free Full Text]
    13. 13
  13. Fullner, K. J., and J. J. Mekalanos. 2000. In vivo covalent cross-linking of cellular actin by the Vibrio cholerae RTX toxin. EMBO J. 19:5315-5323.[CrossRef][Medline]
    14. 14
  14. Gentschev, I., and W. Goebel. 1992. Topological and functional studies on HlyB of Escherichia coli. Mol. Gen. Genet. 232:40-48.[CrossRef][Medline]
    15. 15
  15. Gray, M. C., G. M. Donato, F. R. Jones, T. Kim, and E. L. Hewlett. 2004. Newly secreted adenylate cyclase toxin is responsible for intoxication of target cells by Bordetella pertussis. Mol. Microbiol. 53:1709-1719.[CrossRef][Medline]
    16. 16
  16. Haugen, S. P., M. B. Berkmen, W. Ross, T. Gaal, C. Ward, and R. L. Gourse. 2006. rRNA promoter regulation by nonoptimal binding of sigma region 1.2: an additional recognition element for RNA polymerase. Cell 125:1069-1082.[CrossRef][Medline]
    17. 17
  17. Heidelberg, J. F., J. A. Eisen, W. C. Nelson, R. A. Clayton, M. L. Gwinn, R. J. Dodson, D. H. Haft, E. K. Hickey, J. D. Peterson, L. Umayam, S. R. Gill, K. E. Nelson, T. D. Read, H. Tettelin, D. Richardson, M. D. Ermolaeva, J. Vamathevan, S. Bass, H. Qin, I. Dragoi, P. Sellers, L. McDonald, T. Utterback, R. D. Fleishmann, W. C. Nierman, O. White, S. L. Salzberg, H. O. Smith, R. R. Colwell, J. J. Mekalanos, J. C. Venter, and C. M. Fraser. 2000. DNA sequence of both chromosomes of the cholera pathogen Vibrio cholerae. Nature 406:477-483.[CrossRef][Medline]
    18. 18
  18. Heussen, C., and E. B. Dowdle. 1980. Electrophoretic analysis of plasminogen activators in polyacrylamide gels containing sodium dodecyl sulfate and copolymerized substrates. Anal. Biochem. 102:196-202.[CrossRef][Medline]
    19. 19
  19. Holland, I. B., L. Schmitt, and J. Young. 2005. Type 1 protein secretion in bacteria, the ABC-transporter dependent pathway. Mol. Membr. Biol. 22:29-39.[Medline]
    20. 20
  20. Joelsson, A., Z. Liu, and J. Zhu. 2006. Genetic and phenotypic diversity of quorum-sensing systems in clinical and environmental isolates of Vibrio cholerae. Infect. Immun. 74:1141-1147.[Abstract/Free Full Text]
    21. 21
  21. Koronakis, V., C. Hughes, and E. Koronakis. 1991. Energetically distinct early and late stages of HlyB/HlyD-dependent secretion across both Escherichia coli membranes. EMBO J. 10:3263-3272.[Medline]
    22. 22
  22. Lenz, D. H., M. B. Miller, J. Zhu, R. V. Kulkarni, and B. L. Bassler. 2005. CsrA and three redundant small RNAs regulate quorum sensing in Vibrio cholerae. Mol. Microbiol. 58:1186-1202.[CrossRef][Medline]
    23. 23
  23. Lenz, D. H., K. C. Mok, B. N. Lilley, R. V. Kulkarni, N. S. Wingreen, and B. L. Bassler. 2004. The small RNA chaperone Hfq and multiple small RNAs control quorum sensing in Vibrio harveyi and Vibrio cholerae. Cell 118:69-82.[CrossRef][Medline]
    24. 24
  24. Levine, M. M., J. B. Kaper, D. Herrington, J. Ketley, G. Losonsky, C. O. Tacket, B. Tall, and S. Cryz. 1988. Safety, immunogenicity, and efficacy of recombinant live oral cholera vaccines, CVD 103 and CVD 103-HgR. Lancet ii:467-470.
    25. 25
  25. Lin, W., K. J. Fullner, R. Clayton, J. A. Sexton, M. B. Rogers, K. E. Calia, S. B. Calderwood, C. Fraser, and J. J. Mekalanos. 1999. Identification of a Vibrio cholerae RTX toxin gene cluster that is tightly linked to the cholera toxin prophage. Proc. Natl. Acad. Sci. USA 96:1071-1076.[Abstract/Free Full Text]
    26. 26
  26. Marsh, J. W., and R. K. Taylor. 1998. Identification of the Vibrio cholerae type 4 prepilin peptidase required for cholera toxin secretion and pilus formation. Mol. Microbiol. 29:1481-1492.[CrossRef][Medline]
    27. 27
  27. Mekalanos, J. J. 1983. Duplication and amplification of toxin genes in Vibrio cholerae. Cell 35:253-263.[CrossRef][Medline]
    28. 28
  28. Mel, S. F., K. J. Fullner, S. Wimer-Mackin, W. I. Lencer, and J. J. Mekalanos. 2000. Association of protease activity in Vibrio cholerae vaccine strains with decreases in transcellular epithelial resistance of polarized T84 intestinal epithelial cells. Infect. Immun. 68:6487-6492.[Abstract/Free Full Text]
    29. 29
  29. Miller, J. H. 1972. Experiments in molecular genetics. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY.
    30. 30
  30. Miller, M. B., and B. L. Bassler. 2001. Quorum sensing in bacteria. Annu. Rev. Microbiol. 55:165-199.[CrossRef][Medline]
    31. 31
  31. Miller, M. B., K. Skorupski, D. H. Lenz, R. K. Taylor, and B. L. Bassler. 2002. Parallel quorum sensing systems converge to regulate virulence in Vibrio cholerae. Cell 110:303-314.[CrossRef][Medline]
    32. 32
  32. Murray, H. D., D. A. Schneider, and R. L. Gourse. 2003. Control of rRNA expression by small molecules is dynamic and nonredundant. Mol. Cell. 12:125-134.[CrossRef][Medline]
    33. 33
  33. Oropeza-Wekerle, R. L., E. Müller, P. Kern, R. Meyermann, and W. Goebel. 1989. Synthesis, inactivation, and localization of extracellular and intracellular Escherichia coli hemolysins. J. Bacteriol. 171:2783-2788.[Abstract/Free Full Text]
    34. 34
  34. Sack, D. A., R. B. Sack, G. B. Nair, and A. K. Siddique. 2004. Cholera. Lancet. 363:223-233.[CrossRef][Medline]
    35. 35
  35. Sambrook, J., and D. Russell. 2001. Molecular cloning: a laboratory manual, 3rd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.
    36. 36
  36. Sheahan, K. L., C. L. Cordero, and K. J. Satchell. 2004. Identification of a domain within the multifunctional Vibrio cholerae RTX toxin that covalently cross-links actin. Proc. Natl. Acad. Sci. USA 101:9798-9803.[Abstract/Free Full Text]
    37. 37
  37. Skorupski, K., and R. K. Taylor. 1996. Positive selection vectors for allelic exchange. Gene 169:47-52.[CrossRef][Medline]
    38. 38
  38. Tomarelli, R. M., J. Charney, and M. L. Harding. 1949. The use of azoallbumin as a substrate in colorimetric determination of peptic and tryptic activity. J. Lab. Clin. Med. 34:428-433.[Medline]
    39. 39
  39. Travers, A. A. 1980. Promoter sequence for stringent control of bacterial ribonucleic acid synthesis. J. Bacteriol. 141:973-976.[Abstract/Free Full Text]
    40. 40
  40. Vance, R. E., J. Zhu, and J. J. Mekalanos. 2003. A constitutively active variant of the quorum-sensing regulator LuxO affects protease production and biofilm formation in Vibrio cholerae. Infect. Immun. 71:2571-2576.[Abstract/Free Full Text]
    41. 41
  41. Welch, R. A. 2001. RTX toxin structure and function: a story of numerous anomalies and few analogies in toxin biology. Curr. Top. Microbiol. Immunol. 257:85-111.[Medline]
    42. 42
  42. Yildiz, F. H., X. S. Liu, A. Heydorn, and G. K. Schoolnik. 2004. Molecular analysis of rugosity in a Vibrio cholerae O1 El Tor phase variant. Mol. Microbiol. 53:497-515.[CrossRef][Medline]
    43. 43
  43. Zhu, J., and J. J. Mekalanos. 2003. Quorum sensing-dependent biofilms enhance colonization in Vibrio cholerae. Dev. Cell 5:647-656.[CrossRef][Medline]
    44. 44
  44. Zhu, J., M. B. Miller, R. E. Vance, M. Dziejman, B. L. Bassler, and J. J. Mekalanos. 2002. Quorum-sensing regulators control virulence gene expression in Vibrio cholerae. Proc. Natl. Acad. Sci. USA 99:3129-3134.[Abstract/Free Full Text]

Journal of Bacteriology, March 2007, p. 1827-1835, Vol. 189, No. 5
0021-9193/07/$08.00+0     doi:10.1128/JB.01766-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • Lee, B. C., Lee, J. H., Kim, M. W., Kim, B. S., Oh, M. H., Kim, K.-S., Kim, T. S., Choi, S. H. (2008). Vibrio vulnificus rtxE Is Important for Virulence, and Its Expression Is Induced by Exposure to Host Cells. Infect. Immun. 76: 1509-1517 [Abstract] [Full Text]  
  • Olivier, V., Haines, G. K. III, Tan, Y., Satchell, K. J. F. (2007). Hemolysin and the Multifunctional Autoprocessing RTX Toxin Are Virulence Factors during Intestinal Infection of Mice with Vibrio cholerae El Tor O1 Strains. Infect. Immun. 75: 5035-5042 [Abstract] [Full Text]  

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Boardman, B. K.
Right arrow Articles by Fullner Satchell, K. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Boardman, B. K.
Right arrow Articles by Fullner Satchell, K. J.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»