Transcriptional Regulation of opaR, qrr2–4 and aphA by the Master Quorum-Sensing Regulator OpaR in Vibrio parahaemolyticus

Abstract

Vibrio parahaemolyticus is a leading cause of infectious diarrhea and enterogastritis via the fecal-oral route. V. harveyi is a pathogen of fishes and invertebrates, and has been used as a model for quorum sensing (QS) studies. LuxR is the master QS regulator (MQSR) of V. harveyi, and LuxR-dependent expression of its own gene, qrr2-4 and aphA have been established in V. harveyi. Molecular regulation of target genes by the V. parahaemolyticus MQSR OpaR is still poorly understood.
The bioinformatics analysis indicated that V. parahaemolyticus OpaR, V. harveyi LuxR, V. vulnificu SmcR, and V. alginolyticus ValR were extremely conserved, and that these four MQSRs appeared to recognize the same conserved cis-acting signals, which was represented by the consensus constructs manifesting as a position frequency matrix and as a 20 bp box, within their target promoters. The MQSR box-like sequences were found within the upstream DNA regions of opaR, qrr2-4 and aphA in V. parahaemolyticus, and the direct transcriptional regulation of these target genes by OpaR were further confirmed by multiple biochemical experiments including primer extension assay, gel mobility shift assay, and DNase I footprinting analysis. Translation and transcription starts, core promoter elements for sigma factor recognition, Shine-Dalgarno sequences for ribosome recognition, and OpaR-binding sites were determined for the five target genes of OpaR, which gave a structural map of the OpaR-dependent promoters. Further computational promoter analysis indicated the above regulatory circuits were shared by several other closely related Vibrios but with slight exceptions.
This study gave a comprehensive computational and characterization of the direct transcriptional regulation of five target genes, opaR, qrr2-4 and ahpA, by OpaR in V. parahaemolyticus. These characterized regulatory circuits were conserved in V. harveyi and V. parahaemolyticus.

Full text

Transcriptional Regulation of
opaR
,
qrr2–4
and
aphA
by
the Master Quorum-Sensing Regulator OpaR in
Vibrio
parahaemolyticus
Yiquan Zhang
1.
, Yefeng Qiu
2.
, Yafang Tan
1
, Zhaobiao Guo
1
, Ruifu Yang
1
, Dongsheng Zhou
1
*
1State Key Laboratory of Pathogen and Biosecurity, Beijing Institute of Microbiology and Epidemiology, Beijing, China, 2Laboratory Animal Center, Academy of Military
Medical Sciences, Beijing, China
Abstract
Background:
Vibrio parahaemolyticus is a leading cause of infectious diarrhea and enterogastritis via the fecal-oral route. V.
harveyi is a pathogen of fishes and invertebrates, and has been used as a model for quorum sensing (QS) studies. LuxR is the
master QS regulator (MQSR) of V. harveyi, and LuxR-dependent expression of its own gene, qrr2–4 and aphA have been
established in V. harveyi. Molecular regulation of target genes by the V. parahaemolyticus MQSR OpaR is still poorly
understood.
Methodology/Principal Findings:
The bioinformatics analysis indicated that V. parahaemolyticus OpaR, V. harveyi LuxR, V.
vulnificu SmcR, and V. alginolyticus ValR were extremely conserved, and that these four MQSRs appeared to recognize the
same conserved cis-acting signals, which was represented by the consensus constructs manifesting as a position frequency
matrix and as a 20 bp box, within their target promoters. The MQSR box-like sequences were found within the upstream
DNA regions of opaR,qrr2–4 and aphA in V. parahaemolyticus, and the direct transcriptional regulation of these target genes
by OpaR were further confirmed by multiple biochemical experiments including primer extension assay, gel mobility shift
assay, and DNase I footprinting analysis. Translation and transcription starts, core promoter elements for sigma factor
recognition, Shine-Dalgarno sequences for ribosome recognition, and OpaR-binding sites were determined for the five
target genes of OpaR, which gave a structural map of the OpaR-dependent promoters. Further computational promoter
analysis indicated the above regulatory circuits were shared by several other closely related Vibrios but with slight
exceptions.
Conclusions/Significance:
This study gave a comprehensive computational and characterization of the direct
transcriptional regulation of five target genes, opaR,qrr2–4 and ahpA, by OpaR in V. parahaemolyticus. These characterized
regulatory circuits were conserved in V. harveyi and V. parahaemolyticus.
Citation: Zhang Y, Qiu Y, Tan Y, Guo Z, Yang R, et al. (2012) Transcriptional Regulation of opaR,qrr2–4 and aphA by the Master Quorum-Sensing Regulator OpaR
in Vibrio parahaemolyticus. PLoS ONE 7(4): e34622. doi:10.1371/journal.pone.0034622
Editor: Partha Mukhopadhyay, National Institutes of Health, United States of America
Received December 14, 2011; Accepted March 2, 2012; Published April 10, 2012
Copyright: ß2012 Zhang et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: Financial support was provided by the National Natural Science Foundation of China (31170127, and 30871370), and by the National Basic Research
Program of China (2009CB522604). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: dongshengzhou1977@gmail.com
.These authors contributed equally to this work.
Introduction
Bacterial quorum sensing (QS) is a process of population
density-dependent cell-to-cell communication through synthesiz-
ing, releasing, and detecting signal molecules called autoinducers.
The QS systems are widely distributed in Vibrio species that are
natural inhabitants in seawater, boundary regions between sea and
river, and aquatic products. V. harveyi (a pathogen of fishes and
invertebrates) and V. cholerae (the causative agent of the disease
cholera) have been used as models for QS studies (summarized or
characterized in [1,2,3,4,5,6,7,8,9,10,11,12,13]).
V. harveyi uses three distinct autoinducers, harveyi autoinducer 1
(HAI-1), autoinducer 2 (AI-2), and cholerae autoinducer 1 (CAI-
1), which are synthesized by the autoinducer synthases LuxM,
LuxS, and CqsA, respectively (Fig. 1). They bind to the
membrane-anchoring receptor proteins, LuxN, LuxP/LuxQ,
and CqsS, respectively, at the cell surface. V. cholerae uses two
known autoinducers CAI-1 and AI-2, rather than HAI-1, since the
orthologs of LuxN and LuxM are essentially absent from this
bacterium. The association of autoinducers and their receptor
proteins triggers a common phosphorylation/dephosphorylation
signal transduction cascade involving LuxU and LuxO.
At low cell density (LCD) (Fig. 1), the autoinducers are absent or
at low concentrations, and the receptors autophosphorylate and
then transfer phosphate to the phosphorelay protein LuxU that in
turn shuttles phosphate to the transcriptional factor LuxO. The
phosphorylated LuxO (LuxO-P) in combination with the sigma
factor s
54
activates the transcription of the genes encoding
regulatory small RNAs (sRNAs), Qrr1–4. The Qrr sRNAs
accompanying with the RNA-binding protein Hfq in turns inhibit
the translation of the mRNA of the master QS regulator (MQSR),
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e.g., LuxR in V. harveyi, and HapR in V. cholerae, which leads to the
ceased production of MQSR. The over-production of Qrr sRNAs
and LuxO-P triggers three feedback regulatory loops: i) LuxO-P
represses the transcription of its own gene, ii) Qrr sRNAs inhibits
the translation of luxO, and iii) Qrr sRNAs repress the translation
of luxMN encoding LuxM and its cognate receptor LuxM; these
feedbacks will contribute to control the Qrr levels within
physiological states, since the qrr expression requires the signal
transduction among LuxO, LuxM, and LuxN.
At high cell densities (HCD) (Fig. 1), the accumulated
autoinducers bind to their receptors, and convert the receptors
to phosphatases, thereby reversing the phosphate flow and
triggering the dephosphorylation of LuxU and LuxO. Dephos-
phorylated LuxO cannot activate the qrr transcription. Existing
Qrr sRNAs are rapidly turned over, as Hfq-dependent sRNAs are
degraded stoichiometrically with their target mRNAs. In the
absence of the Qrr sRNAs, the MQSR mRNA is translated, and
MQSR is produced abundantly to act as either a transcriptional
activator or a repressor for its target genes. Overproduced MQSR
will feed back to repress its own transcription.
MQSR is able to activate the transcription of qrr genes, and this
MQSR-qrr feedback loop (Fig. 1) in turns leads to the rapid down-
regulation of MQSR gene. The efficient stimulation of qrr genes
requires the simultaneous presence of LuxO-P (abundant only at
LCD) and MQSR (abundant only at HCD). LuxO-P and MQSR
are thought to be simultaneously present immediately following
the switch from HCD to LCD. Therefore, the above negative
feedback loop dramatically accelerates the transition HCD to
LCD, but it has no effect on the QS behaviors at steady-state LCD
or HCD.
The above five negative feedback loops control the integration
of multiple signals, and maintain the signal transmission fidelity of
QS through affecting the input-output dynamic range of signal
transmission and modulating the noise in the output [13].
V. parahaemolyticus is a leading cause of infectious diarrhea and
enterogastritis via the fecal-oral route [14]. Human infections
occur mainly due to the ingestion of this pathogen in raw or
undercooked seafood. V. harveyi,V. parahaemolyticus, and six
additional closely related species (V. alginolyticus,V. campbellii,V.
rotiferianus,V. natrigens,V. mytili, and V. azureus) constitute the
Harveyi clade that is a subset of the Vibrio core group [15]. All the
components of the V. harveyi QS system can be annotated to be
intact in the genome V. parahaemolyticus [16]. Thus, the signal
transduction cascades of QS should be conserved in V. harveyi and
V. parahaemolyticus (Fig. 1).
The V. parahaemolyticus OpaR [17] is the ortholog of the V. harveyi
LuxR. The opaR gene (VP2516) [16] is composed of an open
reading frame (ORF) containing 615 nucleotides with a G+C
content of 44.55%, and it encodes a deduced protein of 204 amino
acids (a.a.) with a calculated molecular mass of 23634.01 Da and
with an isolectric point of 5.96. Regulation of target genes by
OpaR in V. parahaemolyticus is still poorly understood. In the present
work, the consensus constructs were built to represent the
conserved cis-acting signals recognized by the four extremely
conserved MQSR proteins, V. parahaemolyticus OpaR, V. harveyi
LuxR, V. vulnificus SmcR, and V. alginolyticus ValR, which was
followed by a comprehensive molecular characterization of the
transcriptional regulation of five target genes, opaR,qrr2–4 and
ahpA, by OpaR in V. parahaemolyticus.
Figure 1. Quorum sensing systems in
V. harveyi
/
V. parahaemolyticus
.The mode for signal transduction during QS in V. harveyi has been
described in the text. The feedback regulatory loops are shown with dotted lines. Since all the components of V. harveyi quorum sensing appears to
be intact in the V. parahaemolyticus genome [16], the QS signal transduction cascades should be conserved in V. harveyi and V. parahaemolyticus.
doi:10.1371/journal.pone.0034622.g001
Regulation of opaR,qrr2–4 and aphA by OpaR
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Materials and Methods
Bacterial strains
The wild-type V. parahaemolyticus strain RIMD 2210633 (WT) is
a pandemic O3:K6 strain isolated from a patient with traveler’s
diarrhea in Japan in 1996 [16]. The null opaR mutant derived from
WT and the corresponding complemented mutant are described
below. For the common bacterial growth and maintenance,
bacteria were cultured in Luria-Bertani (LB) broth or LB agar with
addition of 2% NaCl at 37uC, and chloramphenicol was added at
5mg/ml when needed. For the longtime storage, bacteria were
stored in Difco
TM
Marine (MR) broth 2216 (BD Bioscience) with
addition of 30% glycerol at 285uC.
Construction of the opaR null mutant
The entire coding region of opaR was deleted from RIMD
2210633 to generate the opaR null mutant strain DopaR, using the
suicide plasmid pDS132 [18] by introducing homologous
recombination as previously described [19,20]. Briefly, the 414
and 457 bp DNA regions upstream and downstream of opaR,
respectively, were amplified by PCR, purified, and used as the
templates to create a 871 bp deletion construct, through PCR,
which was subsequently inserted between the PstI and SphI sites of
pDS132. This generated a recombinant vector that contained the
deletion construct and the sacB gene conferring sensitivity to
sucrose. All the primers used in the present work were listed in
Table 1. Upon verification by DNA sequencing, the recombinant
vector was introduced into Escherichia coli S17-1(pir), and then
transferred into RIMD 2210633 by conjugation. The mutant
strain was selected using resistance to 10% sucrose and sensitivity
to chloramphenicol, and further verified by PCR.
Complementation of DopaR
For complementation of the DopaR mutant, a PCR-generated
DNA fragment containing the opaR coding region together with its
promoter region (539 bp DNA region upstream of the coding
sequence) and transcriptional terminator region (327 bp DNA
region downstream) were cloned between the SalI and SphI sites of
the vector pBRMob (kindly proved by Prof. Hin-chung Wong
from Taiwan Soochow University) which is the ligation product of
a 3219 bp fragment (containing the RP4 mob DNA region for
plasmid mobilization) from pDS132 digested with HindIII, and the
HindIII-digested plasmid pBR328 (harboring a chloramphenicol
resistance gene) [21]. The recombinant plasmid, verified by DNA
sequencing, was subsequently introduced into DopaR, yielding the
complemented mutant strain C-opaR.
Bacterial growth and RNA isolation
The glyceric stock of bacterial cells was inoculated into 5 ml of
the MR broth for growing at 30uC with shaking at 200 rpm for
24 h. The cell culture was 40-fold diluted with the PBS buffer
(pH 7.2), and then 150 ml of the diluted cells were spread onto a
HI plate [2.5% Bacto heart infusion (BD Bioscience), and 1.5%
bacteriological grade agar] with a diameter of 5 cm. After 8 h of
growth at 30uC, cells were harvested from the plate by adding the
mixture of 1.5 ml of RNAprotect (Qiagen) and 0.5 ml of PBS.
Bacterial cultivations were done at least in triplicate (at least three
biological replicates) for each strain.
Total bacterial RNAs were extracted using the TRIzol Reagent
(Invitrogen) [22,23,24]. RNA quality was monitored by agarose
gel electrophoresis, and RNA quantity was determined by
spectrophotometry.
Primer extension assay
For the primer extension assay [22,23,24], an oligonucleotide
primer complementary to a portion of the RNA transcript of each
indicated gene was employed to synthesize cDNAs from the RNA
templates. About 10 mg of the total RNA from each strain was
annealed with 1 pmol of [c-
32
P] end-labeled reverse primer using
a Primer Extension System (Promega) according to the manufac-
turer’s instructions. The same labeled primer was also used for
sequencing with the fmolHDNA Cycle Sequencing System
(Promega). The primer extension products and sequencing
materials were concentrated and analyzed in a 6% polyacryl-
amide/8 M urea gel. The result was detected by autoradiography
(Kodak film).
Preparation of purified OpaR protein
Preparation of the purified OpaR protein was performed as
previously described [22,23,24]. The entire coding region of the
opaR gene of strain RIMD 2210633 was directionally cloned
between the BamHI and HindIII sites of plasmid pET28a
(Novagen). The recombinant plasmid encoding the 66His-tagged
OpaR protein (His-OpaR) was transformed into Escherichia coli
BL21lDE3 cells. Expression of His-OpaR was induced by the
addition of 1 mM IPTG (isopropyl-b-D-thiogalactoside). The
overproduced protein was purified under native conditions using
an Ni-NTA Agarose Column (Qiagen). The purified protein was
concentrated with the Amicon Ultra-15 centrifugal filter device
(Millipore), and the protein purity was verified by SDS-PAGE.
Gel mobility shift assay (EMSA)
The 300 to 600 bp promoter-proximal region of each indicated
gene was amplified by PCR. For EMSA [22,23,24], the 59ends of
DNA were labeled using [c-
32
P] ATP and T4 polynucleotide
kinase. DNA binding was performed in a 10 ml reaction volume
containing binding buffer [1 mM MgCl
2
, 0.5 mM EDTA,
0.5 mM DTT, 50 mM NaCl, 10 mM Tris-HCl (pH 7.5) and
0.05 mg/ml poly-(dI-dC)], labeled DNA (1000 to 2000 c.p.m/ml),
and increasing amounts of the His-OpaR protein. Three controls
were included in each EMSA experiment: 1) cold probe as specific
DNA competitor (the same promoter-proximal DNA region
unlabeled), 2) negative probe as nonspecific DNA competitor
(the unlabeled coding region of the 16S rRNA gene), and 3)
nonspecific protein competitor [rabbit anti-F1-protein polyclonal
antibodies]. The F1 protein is the protective antigen from Yersinia
pestis [25]. After incubation at room temperature for 30 min, the
products were loaded onto a native 4% (w/v) polyacrylamide gel,
and electrophoresed in 0.56TBE buffer for about 50 min at
220 V. Radioactive species were detected by autoradiography
after exposure to Kodak film at 270uC.
DNase I footprinting
For DNase I footprinting [22,23,24], the 250 to 600 bp
promoter-proximal DNA regions with a single
32
P-labeled end
were PCR amplified with either the sense or antisense primer
being end-labeled. The PCR products were purified using the
QiaQuick columns (Qiagen). Increasing amounts of His-OpaR
were incubated with the purified, labeled DNA fragment (2 to
5 pmol) for 30 min at room temperature, in a final 10 ml reaction
volume containing the binding buffer used in EMSA. Before DNA
digestion, 10 mlofCa
2+
/Mg
2+
solution (5 mM CaCl
2
and 10 mM
MgCl
2
) was added, followed by incubation for 1 min at room
Regulation of opaR,qrr2–4 and aphA by OpaR
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temperature. The optimized RQ1 RNase-Free DNase I (Promega)
was then added to the reaction mixture, and the mixture was
incubated at room temperature for 40 to 90 s. The reaction was
quenched by adding 9 ml of stop solution (200 mM NaCl, 30 mM
EDTA, and 1% SDS), followed by incubation for 1 min at room
temperature. The partially digested DNA samples were extracted
with phenol/chloroform, precipitated with ethanol, and analyzed
in 6% polyacrylamide/8 M urea gel. Protected regions were
identified by comparison with the sequence ladders. For
sequencing, we used the fmolHDNA Cycle Sequencing System
(Promega). The templates for sequencing were the same as the
DNA fragments of DNase I footprinting assays. Radioactive
species were detected as previously described.
Computational promoter analysis
The 300 bp promoter regions upstream of the start codon of
each indicated gene were retrieved with the ‘retrieve-seq’ program
[26]. Known or predicted binding sites of OpaR and its orthologs
were collected and aligned to generate the position frequency
matrix (PFM) by using the ‘matrices-consensus’ tool [26].The
sequence logo representation of the above binding sites was
generated by the WebLogo tool [27]. The ‘matrices-paster’ tool [26]
was used to match the PFM within the promoter-proximal DNA
regions.
Results
Phylogeny of OpaR and its orthologs
The OpaR regulator shares high identity ($70%) in a.a.
sequences with the orthologous MQSRs in other six Vibrio species
tested (V. alginolyticus ValR, V. harveyi LuxR, V. vulnificus SmcR, V.
tubiashii VtpR, V. anguillarum VanT, and V. cholerae HapR). A
phylogenetic tree (Fig. 2) was constructed from the aligned a.a.
sequences of the above seven orthologous MQSR proteins, with
an additional regulator LitR from V. fischeri [28] as the outgroup
(LitR has about 60% identity to the above seven MQSRs; all these
eight proteins are belonged to the TetR-family DNA-binding
regulators), which revealed that LuxR, OpaR, SmcR, and ValR
constituted the most closely related group (.92% identity between
each other).
The MQSR consensus
Since the four DNA-binding regulatory proteins LuxR, OpaR,
SmcR, and ValR were extremely conserved, they should recognize
the same conserved signals within their target promoters in V.
harveyi,V. parahaemolyticus,V. vulnificus, and V. alginolyticus. Known or
predicted binding sites of LuxR, OpaR, SmcR, and ValR (Table 2)
were collected, and then aligned to generate the MQSR consensus
that manifested as a PFM (in which each row and column
represents a position and a nucleotide, respectively) and as a 20 bp
Table 1. Oligonucleotide primers used in this study.
Target Primers (forward/reverse, 59-39)
Construction of mutant
opaR GTGACTGCAGACTGCCTTGGTAACGCTCTG/GTTCGTGTTCAAATCTGAGCTATCCATTTTCCTTGCCATTTG
CAAATGGCAAGGAAAATGGATAGCTCAGATTTGAACACGAAC/GTGAGCATGCATGGGCTGCATCAGGTCG
GTGACTGCAGACTGCCTTGGTAACGCTCTG/GTGAGCATGCATGGGCTGCATCAGGTCG
Complementation of mutant
opaR GCGGGATCCTCCATCGTGTTGCCGTAGC/GCGAAGCTTGCGAAAGCAGAAGGCATCAAG
Protein expression
opaR AGCGGGATCCATGGACTCAATTGCAAAGAG/AGCGAAGCTTTTAGTGTTCGCGATTGTAG
EMSA
opaR TGTGGGTTGAGGTAGGTCG/GCCTAGTTCTAGGTCTCTTTGC
qrr2 AGTGGTTGCTTATGAATC/GGTCGAGAAGTATTATGC
qrr3 GGATAAGTTCAAATTGGATC/GTGGTTTCTGTGACATAC
qrr4 AACCGTGAAATCCATTTAC/CGACGCATTATTAACCAG
aphA AACTTCCAACCACATAATTGCG/GGCTGGAGCAGGTATGATTG
DNase I footprinting
opaR AGTGGGTTGAAAGTCACATCC/GCCTAGTTCTAGGTCTCTTTGC
qrr2 AGTGGTTGCTTATGAATC/GGTCGAGAAGTATTATGC
qrr3 GGATAAGTTCAAATTGGATC/GTGGTTTCTGTGACATAC
qrr4 AACCGTGAAATCCATTTAC/CGACGCATTATTAACCAG
aphA AACTTCCAACCACATAATTGCG/GGCTGGAGCAGGTATGATTG
Primer extension
opaR /ATCCATTTTCCTTGCCATTTG
qrr2 /TTATTGTGAACAATCTATAT
qrr3 /AATCAAGTTCACTAACAAC
qrr4 /ATATACTTGTGAACAATGTG
aphA /GCTCTTACTGGCGCTTGAG
doi:10.1371/journal.pone.0034622.t001
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invert-repeat sequence TATTGATAAA-TTTATCAATA termed
as the MQSR box (Fig. 3).
LuxR-dependent expression of its own gene [29], qrr2–4 [7],
and aphA [30] have been established in V. harveyi. The presence of
MQSR box-like sequences within the upstream DNA regions of
the corresponding target genes in V. parahaemolyticus, as revealed by
the computational promoter analysis (Table 2), indicated that the
above regulatory cascades were conservatively controlled by the
LuxR ortholog OpaR in V. parahaemolyticus, which were further
validated by the following biochemical experiments.
Mutation and complementation of opaR
Real-time RT-PCR experiments were performed to assess the
relative mRNA levels of opaR in WT, DopaR, and C-opaR; the opaR
transcript was lacking in DopaR, but was restored in C-opaR relative
to WT (data not shown), indicating the successful mutation of opaR
and the complementation of the opaR mRNA level.
As determined by several distinct methods (see below), the ahpA
gene was negatively regulated by OpaR. To test whether the opaR
mutation had the polar effect, the primer extension assays were
conducted to detect the yield of the primer extension product of
ahpA that represented the ahpA mRNA levels in WT, DopaR, and
C-opaR. Herein, the ahpA mRNA level was significantly enhanced
in DopaR relative to WT, while no obvious change in the ahpA
transcription was observed between WT and C-opaR (Fig. S1). This
analysis confirmed that the detecting enhanced transcription of
ahpA in DopaR was due to the opaR mutation rather than a polar
effect.
Figure 2. Phylogenetic tree of OpaR and its orthologs. Protein sequences were derived from V. alginolyticus ZJ-51 [60], V. parahaemolyticus
RIMD 2210633 [16], V. harveyi ATCC BAA-1116 [61], V. vulnificus YJ016 [62], V. tubiashii RE22 [63], V. anguillarum 775 [64], V. cholera N16961 [65], and V.
fischeri MJ11 [66]. The a.a. sequences were aligned by the CLUSTALW [67] web server at http://align.genome.jp/. The aligned sequences were then
used to construct an unrooted neighbor-joining tree using the MEGA version 5.0 [68] with a bootstrap iteration number of 1000. Shown on the
branch points of phylogenic tree were the bootstrap values (%).
doi:10.1371/journal.pone.0034622.g002
Figure 3. MQSR consensus constructs. (a) The sequence logo representation of the binding sites of OpaR and its orthologs (Table 2) was
generated by the WebLogo tool [27]. The 20-bp consensus box TATTGATAAA-TTTATCAATA was annotated as an inverted repeat sequence. (b) A
position frequency matrix describes the alignment of the binding sites, and denotes the frequency of each nucleotide at each position.
doi:10.1371/journal.pone.0034622.g003
Regulation of opaR,qrr2–4 and aphA by OpaR
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Growth of WT and DopaR
The growth curves of WT and DopaR grown at 30uC in the MR
broth or in the HI broth were determined (Fig. 4). The two strains
showed indistinguishable growth rates in each of the media. Thus,
the opaR mutation had no effect on the bacterial in vitro growth.
For the following molecular regulation experiments, bacteria
were pre-cultivated in the MR broth, spread onto grown on the HI
agar plates for further growth, and harvested after an 8 h
incubation at 30uC [at this status, the dense bacterial lawns (i.e.
HCD) were observed on the agar]. It was thought that, unlike the
liquid cultivation for which the autoinducers would disperse into
the liquid media, the cultivation on solid medium would enable
the enrichment of autoinducer molecules within the bacterial
lawns with little dispersal into the agar.
Negative auto-regulation of OpaR
The primer extension experiments (Fig. 5a) were conducted to
compare the yields of primer extension product of opaR in WT and
DopaR. The primer extension assay detected a single transcription
start site located at 74 bp upstream of opaR; therefore, a single
promoter was transcribed for opaR under the growth condition
tested. In addition, the opaR promoter activity was under the
negative control of OpaR. A 334 bp promoter-proximal region of
opaR was amplified, radioactively labeled, and subjected to EMSA
with a purified His-OpaR protein (Fig. 5b). The results showed
that His-OpaR was able to bind to this DNA fragment in a dose-
dependent manner in vitro. As further determined by DNase I
footprinting (Fig. 5c), the purified His-OpaR protein protected two
distinct regions upstream of opaR against DNase I digestion in a
dose-dependent manner. These two footprints, located from 70 to
40 bp (site 1) and from 159 to 109 bp (site 2) upstream of opaR,
respectively, were considered as OpaR-binding sites. Taken
together, OpaR is able to recognize the promoter of its own gene
to directly repress its activity in V. parahaemolyticus.
Stimulation of qrr2–4 by OpaR
The primer extension assay (Fig. 6a) defined the transcription
start sites the three sRNA genes qrr2–4, and this assay also
indicated that the promoter activity of all the thee qrr genes was
under the positive control of OpaR. Each of the promoter-
proximal regions of qrr2–4 was subjected to EMSA with the
purified His-OpaR protein (Fig. 6b). The results showed that His-
OpaR was able to bind to each of the three DNA fragments tested
in a dose-dependent manner in vitro. As further determined by
DNase I footprinting (Fig. 6c), His-OpaR protected a single region
within each of the three upstream DNA fragments tested against
Table 2. Known or predicted direct targets of OpaR or its orthologs.
Gene name Gene ID Consensus-like sequence Position
&
Score Reference Regulation
V. harveyi
ATCC BAA-1116
luxR VIBHAR_03459 TAATGACATTACTGTCTATA D-71…-52 9.99 [29] $
AACTATTAAAATAATCAATT D-152…-133 10.93 [29] $
qrr2 VIBHAR_04846 TGATGATTTATTTATCACTT D-165…-146 7.62 [7] #
qrr3 VIBHAR_05322 AGTTAATTAATTCATCATTA D-151…-132 8.02 [7] #
qrr4 VIBHAR_06697 TTCTGATAAATGTATTAGTA D-167…-148 9.34 [7] #
aphA VIBHAR_00046 TATTGAGTATTTTATTAGTT D-281…-262 12.04 [30] $
exsB VIBHAR_01694 TTTTAATAAAAAGATAAGTA D-135…-116 6.45 [31] $
qrgB VIBHAR_00176 TATTGATTGTGAACTCAATA D-98…-79 10.25 [3,30] $
luxC VIBHAR_06244 TACAAATAACATTAATAATT D-275…-256 8.49 [58] #
TATAAATAAATCAAACTATA D-151…-132 8.88 [58] #
argA VIBHAR_03295 AATTGAATAAGAAGACAATA D-64…-45 6.6 [59] #
V. parahaemolyticus
RIMD 2210633
opaR VP2516 TAATGACATTACTGTCTATA D-149…-130 9.99 This study $
AATTATTAAAATAATCAATT D-68…-49 11.29 This study $
qrr2 VPA1623–1624 intergenic GACTAACTCAATTGTTAATA D-167…-148 7.8 This study #
qrr3 VPA1240–1241 intergenic TGTTTATTAATCAATCATTA D-150…-131 8.5 This study #
qrr4 VPA0199–0200 intergenic TGCTGAGAAAGTGATTAGTA D-167…-148 8.16 This study #
aphA VP2762 TATTGAGTATTATGTTAGTT D-279…-260 10.68 This study $
V. vulnificus
YJ016
smcR VV2770 TATTGACATTACTGTTCATT D-158…-139 8.56 Predicted $
AATTATTAAAACAATCAATA D-78…-59 11.33 Predicted $
qrr3 TATAAATAGATTTATTATTA D-191…-172 10.79 Predicted #
aphA VV3005 TATTGAGCATTTTGTTAGTT D-278…-259 10.50 Predicted $
V. alginolyticus
ZJ-51
valR TAATGACATTACTGTATATA D-146…-127 7.60 Predicted $
AACTATTAAAATAATCAATT D-65…-46 10.93 Predicted $
&
, ‘D’ indicates the direct sequence, and minus numbers denote nucleotide positions upstream of genes.
Negative ($) or positive (#) regulation by LuxR or its relevant ortholog.
doi:10.1371/journal.pone.0034622.t002
Regulation of opaR,qrr2–4 and aphA by OpaR
PLoS ONE | www.plosone.org 6 April 2012 | Volume 7 | Issue 4 | e34622

DNase I digestion in a dose-dependent manner. These footprints,
located from 172 to 143 bp, from 154 to 125 bp, and from 204 to
133 bp upstream of qrr2–4, respectively, were considered as
OpaR-binding sites for these three genes. Taken together, OpaR is
able to recognize the promoters of qrr2–4 to activate their activity
in V. parahaemolyticus.
Repression of aphA by OpaR
The primer extension assay (Fig. 7a) detected two closely
neighboring extension products for aphA. Due to the facts that the
shorter extension product might represent the premature stops due
to the difficulty of polymerase in passing difficult sequences, and
that the core promoter elements recognized by sigma factors could
not be identified for the shorter extension product, only the longer
product was chosen for the identification of the transcription start
site that was located at 200 bp upstream of aphA. Therefore, a
single promoter was transcribed for aphA under the growth
condition tested, and its activity was under the negative control of
OpaR. A 541 bp promoter-proximal region of aphA was subjected
to EMSA with the purified His-OpaR protein (Fig. 7b). The results
Figure 4. Growth curves. A two-round design of bacterial seed cultivation was employed: first, the glyceric stock of bacteria was inoculated into
15 ml of the MR or HI broth for growing at 30uC for 24 h with shaking at 200 rpm, and the cell culture was subsequently diluted to an OD
600
value of
about 1.0; second, the resulting culture was then 50-fold diluted into 15 ml of corresponding fresh MR or HI broth, and allowed to grow to reach an
OD
600
value of about 1.2 to 1.4. The bacterial seeds were 50-fold diluted into 15 ml of corresponding fresh MR or HI broth for further cultivation, and
the OD
600
values were monitored for each culture with a 1 h interval. Experiments were done in triplicate.
doi:10.1371/journal.pone.0034622.g004
Figure 5. Repression of its own gene by OpaR. a)Primer extension. An oligonucleotide primer was designed to be complementary to the
RNA transcript of opaR. The primer extension products were analyzed with 8 M urea-6% acrylamide sequencing gel. Lanes C, T, A, and G represent
the Sanger sequencing reactions. The transcription start site of opaR was underlined in the DNA sequence. b)EMSA. The radioactively labeled DNA
fragment from the 300th bp upstream to the 34th bp downstream of opaR was incubated with increasing amounts of purified His-OpaR protein, and
then subjected to 4% (w/v) polyacrylamide gel electrophoresis. The band of free DNA disappeared with increasing amounts of His-OpaR protein, and
a retarded DNA band with decreased mobility turned up, which presumably represented the DNA-OpaR complex. Shown on the lower side of the
figure was the schematic representation of the EMSA design. c)DNase I footprinting. Labeled coding or non-coding DNA probes were incubated
with increasing amounts of purified His-OpaR (Lanes 1, 2, 3, and 4 containing 0, 6, 9, and 12 pmol, respectively), and subjected to DNase I footprinting
assay. Lanes G, A, T, and C represented the Sanger sequencing reactions. The footprint regions were indicated with vertical bars. The negative or
positive numbers indicated the nucleotide positions upstream or downstream of opaR, respectively.
doi:10.1371/journal.pone.0034622.g005
Regulation of opaR,qrr2–4 and aphA by OpaR
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Figure 6. Stimulation
qrr2–4
by OpaR. For primer extension (a1, b1, and c1), an oligonucleotide primer was designed to be complementary to the
RNA transcript of each of qrr2–4. For EMSA (a2, b2, and c2) and DNase I footprinting (a3, b3, and c3), the upstream DNA fragments of qrr2–4 were
radioactively labeled, and then incubated with increasing amounts of purified His-OpaR protein. The experiments were done as described in Fig. 5.
The transcription start sites of qrr2–4 were underlined in the DNA sequence. Lanes G, A, T, and C represented the Sanger sequencing reactions. The
footprint regions were indicated with vertical bars. The negative or positive numbers indicated the nucleotide positions upstream or downstream of
relevant qrr gene, respectively.
doi:10.1371/journal.pone.0034622.g006
Regulation of opaR,qrr2–4 and aphA by OpaR
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showed that His-OpaR was able to bind to the DNA fragment in a
dose-dependent manner in vitro. As further determined by DNase I
footprinting (Fig. 7c), His-OpaR protected a single region from
284 to 255 bp upstream of opaR against DNase I digestion in a
dose-dependent manner. This footprint was considered as the
OpaR-binding site for aphA. Taken together, OpaR is able to
recognize the promoter of aphA to repress its activity in V.
parahaemolyticus.
Discussion
Regulation of biofilm formation and virulence by Vibrio
MQSRs
The V. harveyi LuxR mediates the repression of virulence
determinant type III secretion system 1 (T3SS1) [31] and the
stimulation of bioluminescence encoded by the luxCDABEGH
operon [32]. LuxR binds to the upstream DNA region of
luxCDABEGH, and thus stimulates this operon in a direct manner
[32]; in contrast, LuxR indirectly represses the T3SS1 genes via
directly repressing of exsA, a transcriptional activator of T3SS1,
within the T3SS1 gene loci [31].
The V. cholerae HapR is a repressor of virulence: i) HapR directly
represses the transcription of aphA [33] encoding a regulator [34]
that in turns stimulates the expression of toxin-coregulated pilus;
and ii) HapR inhibits the hemolytic activity at both transcriptional
and posttranslational levels [35] (for the former mechanism, HapR
directly represses the transcription of the hemolysin gene hlyA; for
the later one, HapR directly stimulates the transcription of hapA
encoding a metalloprotease that in turns degrades the HlyA
hemolysin). HapR is also a repressor of biofim formation in V.
cholerae [36]: i) HapR directly inhibits the transcription of vpsT
encoding a transcriptional activator of biofilm formation; and ii)
HapR represses the cellular c-di-GMP levels (c-di-GMP in turns
acts as a posttranscriptional activator of the biofilm formation [37])
through directly modulating the transcription of multiple genes
encoding GGDEF and EAL proteins.
The V. vulnificus SmcR is a repressor of cytotoxicity [38] that is
important for the virulence of V. vulnificus and mainly dependent
on two cytotoxins, RTX (encoded by rtxA1) and cytolysin/
hemolysin (encoded by vvhA) [39,40]. The transcription of rtxA1
and vvhA is repressed by SmcR through directly repressing the
transcription of hlyU [38], an activator of rtxA1 and vvhA [40,41].
SmcR-dependent expression of the metalloprotease gene vvpE is
also found in V. vulnificus [42]; the two regulators cAMP receptor
protein (CRP) and SmcR bind to the upstream region of vvpE in a
juxtapositioned manner, and thus they function synergistically to
coactivate the transcription of vvpE by the RpoS-dependent
promoter at the stationary growth phase [42]. Whether the
SmcR-dependent stimulation of metalloprotease contributes to the
degradation of relevant protein toxins is still unclear in V. vulnificus.
The V. parahaemolyticus OpaR is a repressor of cytotoxicity to
host cells [43], most likely through inhibiting the assembly/
secretion of the cytotoxicity determinant T3SS1 [43,44]. OpaR
appears to repress biofilm formation through directly modulating
the transcription of multiple genes encoding GGDEF and EAL
proteins in pandemic O3:K6 V. parahaemolyticus (data unpublished).
The molecular mechanisms employed by OpaR to regulate
biofilm formation and virulence need to be elucidated.
Biofilm formation can be concisely linked to the bacterial
survive in adverse conditions outside of the host, thus aiding in
bacterial persistence during inter-epidemic seasons [45]. As shown
in V. cholerae [45,46], both intact and dispersed biofilms enhance
the bacterial infectivity upon oral ingestion. Based on the previous
speculations for V. cholerae [36,47,48,49], a model of regulation of
biofilm formation and virulence by QS during intestinal infection
of pathogenic Vibrios is proposed herein: on the initial
colonization (i.e., LCD) of a host, the expression of the MQSRs
was inhibited by the Qrr sRNAs, and thus the expression of the
Figure 7. Repression of
aphA
by OpaR. For primer extension (a), an oligonucleotide primer was designed to be complementary to the RNA
transcript of aphA. For EMSA (b) and DNase I footprinting (c), the DNA fragment from the 380th to 161th bp upstream of aphA was incubated with
increasing amounts of purified His-OpaR protein. The experiments were done as described in Fig. 5. The transcription start site of aphA were
underlined in the DNA sequence. Lanes G, A, T, and C represented the Sanger sequencing reactions. The footprint regions were indicated with
vertical bars.. The minus numbers indicated the nucleotide positions upstream of aphA.
doi:10.1371/journal.pone.0034622.g007
Regulation of opaR,qrr2–4 and aphA by OpaR
PLoS ONE | www.plosone.org 9 April 2012 | Volume 7 | Issue 4 | e34622

biofilm formation and virulence genes occurs, which promotes the
bacterial colonization and infection. When a HCD is reached, the
MQSRs are abundantly produced, and thus inhibit biofilm
formation and virulence in both direct (via control of structural
genes) and indirect (via modulation of regulatory determinants)
manners.
Conserved cis-acting DNA signals recognized by MQSRs
Two consensus constructs, a box and a PFM, (Fig. 3) were built
to represent the conserved cis-acting signals recognized by the four
extremely conserved MQSR proteins, V. harveyi LuxR, V.
parahaemolyticus OpaR, V. vulnificus SmcR, and V. alginolyticus ValR.
These consensus constructs could be also applied to all the other
members of the Harveyi clade in addition to the above four
bacteria.
The 20 bp MQSR box was further annotated as an inverted
repeat sequence. This dyad symmetry structure indicated that
LuxR, OpaR, SmcR, and ValR, like other TetR-type proteins,
bind to cis-acting regulatory DNA as a dimer. The box is a
contiguous oligonucleotide, and thus it presents limited informa-
tion that are originally presented in the MQSR-binding sites.
Representation of the cis-acting regulatory motif with a PFM is
able to give a much more comprehensive description of the uneven
composition in each position, i.e., some nucleotides occurred
much more frequently than others. Thus, the PFM presentedd
here will over-represent the MQSR-binding sites more accurately
than the 20 bp box sequence.
The PFM can be used to statistically predict the presence of
MQSR consensus-like elements within the promoter-proximal
sequences tested, which will generate a weight score for each gene,
and a higher score denoted the higher probability of regulator-
promoter association (Table 2). This assay was applied to the
OpaR regulon members previously determined by microarray
[43], when a frequently-used cutoff number of 7 was set for the
score values, disclosing a set of candidates of direct OpaR targets
(data not shown) in V. parahaemolyticus for further biochemical
validation.
The PFM construct herein is essentially in agreement with those
previously characterized for LuxR in V. harveyi [30] and SmcR in
V. vulnificus [50]. A major difference is that the PFM of this study
was constructed from the cis-acting DNA sequences from four
closed related Vibrios, rather than from the artificial sequences
[30,50]. It was deemed that the PFM herein would enable the
more accurate prediction of novel MQSR box-like sequences.
Autoregulation of MQSRs
Direct transcriptional repression of their own genes have been
established for LuxR [29], HapR [51], VanT [52], and OpaR (this
study). The promoter-proximal regions of valR,opaR,luxR,smcR,
vtpR,vanT, and hapR were aligned in Fig. 8a, in which shown were
translation and transcription starts, 235 and 210 core promoter
elements for s
70
recognition, Shine-Dalgarno (SD) sequences for
ribosome recognition, and MQSR box-like sequences representing
the conserved signals for recognition by OpaR or its orthologs.
This analysis gave a structural map of these auto-repressed
promoters.
Two MQSR box-like sequences, upstream and downstream of
the transcription start site, respectively, were annotated for valR,
opaR,luxR,orsmcR, indicating that two sites were recognized by
the relevant regulatory protein for each target gene. Indeed,
corresponding two binding sites have been experimentally
determined for opaR (this study) or luxR [29]. Only one MQSR
box-like sequence downstream of the transcription start site was
annotated for vtpR,vanT,orhapR, indicating a single site within
each of these promoter-proximal regions were recognized by the
relevant regulatory protein; without no exception, a single
corresponding HapR-binding site has been detected for hapR
[51]. Notably, the MQSR box-like sequences downstream of the
transcription start site were conservatively located within all the
target promoter regions aligned, and the MQSR-promoter DNA
association would block the entry of the RNA polymerase to
repress the transcription of the target genes.
Regulation of Qrr sRNAs genes by MQSRs
All of the three closely related organisms V. harveyi,V.
parahaemolyticus, and V. vulnificus contain qrr1–5, and whereas, the
more distantly related V. cholerae harbors only qrr1–4. Any one of
Qrr1–4 in V. cholerae is perfectly sufficient to repress hapR, and thus,
the four Qrr sRNAs are functionally redundant [2,11]. Qrr1–4 but
not Qrr5 are functional in V. harveyi, and Qrr5 may be an
evolutionary vestige [4]. Unlike in V. cholerae, Qrr1–4 in V. harveyi
function additively to control QS behaviors; these sRNAs function
to translate increasing autoinducer concentrations (following the
transition from LCD to HCD) into a precise gradient of LuxR
protein, and that LuxR in turns induces a gradient of expression its
target genes [4]. Functions of Qrr sRNAs in V. parahaemolyticus and
V. vulnificus need to be elucidated.
As mentioned above, the LuxR- or OpaR-mediated stimulation
of qrr transcription constitutes a negative feedback loop most likely
maintaining the QS behaviors during the transition HCD to LCD
[7,8]. The V. harveyi LuxR directly binds to the upstream DNA
regions of qrr2–4 and stimulates the transcription of qrr2–4, but it
has no regulatory effect on qrr1 and qrr5 [7]. In this work, we
confirmed that V. parahaemolyticus OpaR also bound to the
promoter regions of qrr2–4 to activate their transcription. To the
best of our knowledge, this is the first report of experimentally
determining MQSR-binding sites and transcription starts of qrr
genes. It should be noted the experimental transcription starts of
qrr genes (this study) are 1 bp downstream of the predicted ones
[2].
MQSR box-like sequences were found within the promoter-
proximal regions of qrr2–4 in V. harveyi and V. parahaemolyticus, and
of qrr3 in V. vulnificus (but not qrr2 and qrr4 of V. vulnificus). In
addition, the box elements could not be predicted from qrr1 and
qrr5 of all these three closely related Vibrios. These indicated that
the MQSR-qrr2–4 feedback loop was conversed between V. harveyi
and V. parahaemolyticus. We aligned the promoter-proximal regions
of qrr2–4 (Fig. 8a) from V. harveyi,V. parahaemolyticus, and V.
vulnificus, depicting translation and transcription starts, 224 and
212 core promoter elements for s
45
recognition, and MQSR box-
like sequences (Fig. 8c). Since MQSR box-like sequences are
upstream of the promoter 224 elements, the MQSR-stimulated
qrr promoters may have a class I regulation that depends on the
RNA polymerase asubunit C-terminal domain for function [53].
Unlike in V. harveyi, the V. cholerae HapR stimulates the
transcription of all the qrr1–4 in an indirect manner, since none
of the binding of HapR to the qrr upstream regions can be detected
[8]. As expected, MQSR box-like sequences could not be
predicted from the upstream regions of all of qrr1–4 in V. cholerae.
Regulation of aphA by MQSRs
The AphA regulator is required for auto-repression [33],
intestinal colonization and virulence [34,54], biofilm formation
[55,56] in V. cholerae. In addition, as previously summarized [57],
AphA and LuxR/HapR reciprocally control QS behaviors in V.
harveyi and V. cholerae. At LCD, redundant Qrr sRNAs promote the
AphA translation and meanwhile inhibit the LuxR translation.
AphA further directly represses the luxR transcription, and also
Regulation of opaR,qrr2–4 and aphA by OpaR
PLoS ONE | www.plosone.org 10 April 2012 | Volume 7 | Issue 4 | e34622

feeds back to repress the qrr transcription. At HCD, the cessation
of Qrr sRNA production leads to no production of AphA, and the
LuxR translation occurs. LuxR in turns directly represses the aphA
transcription, and also feeds back to inhibit it own expression.
Thus, AphA acts as a master regulator of QS behaviors at LCD,
and in contrast, LuxR/HapR is the major one operating at HCD;
the reciprocal gradients of AphA and LuxR/HapR are thought to
be established for controlling the gene expression patterns during
the transition between LCD and HCD [57].
The transcription of aphA is directly repressed by LuxR in V.
harveyi [30] and by HapR V. cholerae [33,48], and moreover HapR-
binding site and transcription start have been determined for aphA
in V. cholerae [48]. This study also detected the direct transcrip-
tional repression of aphA by OpaR with the determination of
transcription start and OpaR-binding site for aphA in V.
parahaemolyticus. The alignment of the upstream regions of aphA
from V. harveyi,V. parahaemolyticus,V. vulnificus and V. cholerae
indicated that these four bacteria employed a conserved molecular
mechanism for the repression of aphA by MQSRs, since
transcription starts, 235 and 210 core promoter elements, SD
sequences, and MQSR box-like sequences are conservatively
located upstream of the aphA translation starts.
Supporting Information
Figure S1 Primer extension assay for validation of non-
polar mutation. The opaR null mutant DopaR was generated
from the wild-type (WT) strain RIMD 2210633, and then the
complemented mutant strain C-opaR was constructed. As deter-
mined by several distinct methods (see text), the transcription of
ahpA was under the negative control of OpaR. Herein, an
oligonucleotide primer, which was complementary to the RNA
transcript of ahpA, was employed to detect the primer extension
product that represented the relative mRNA level of ahpA in WT,
DopaR, and C-opaR. The primer extension products were analyzed
with 8 M urea–6% acrylamide sequencing gel. Lanes C, T, A, and
G represented the Sanger sequencing reactions. The transcription
start site (nucleotide C), which was located at 200 bp upstream of
ahpA, was underlined in the DNA sequence. The ahpA mRNA level
was significantly enhanced in DopaR relative to WT, while no
obvious change in the ahpA transcription was observed between
WT and C-opaR, which confirmed that the detecting enhanced
transcription of ahpA in DopaR was due to the opaR mutation rather
than a polar mutation.
(TIF)
Acknowledgments
We thank Professor Mitsuaki Nishibuchi from Kyoto University for kindly
providing the strain RIMD 2210633. The English writing of the
manuscript was polished by EnPapers.
Author Contributions
Conceived and designed the experiments: DZ. Performed the experiments:
YZ YQ YT ZG DZ. Analyzed the data: DZ YZ YQ. Contributed
reagents/materials/analysis tools: DZ YZ YQ. Wrote the paper: DZ YZ
RY.
Figure 8. Organization of promoter DNA regions. DNA sequences were derived from V. alginolyticus ZJ-51 [60], V. parahaemolyticus RIMD
2210633 [16], V. harveyi ATCC BAA-1116 [61], V. vulnificus YJ016 [62], V. tubiashii RE22 [63], V. anguillarum 775 [64], V. cholera N16961 [65], and V.
fischeri MJ11 [66]. Shown were translation and transcription starts, SD sequences, MQSR box-like sequences, and 210/212 and 235/224 core
promoter elements.
doi:10.1371/journal.pone.0034622.g008
Regulation of opaR,qrr2–4 and aphA by OpaR
PLoS ONE | www.plosone.org 11 April 2012 | Volume 7 | Issue 4 | e34622

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