Helicobacter pylori perceives the quorum-sensing
molecule AI-2 as a chemorepellent via the
Bethany A. Rader,13 Christopher Wreden,1Kevin G. Hicks,14
Emily Goers Sweeney,1Karen M. Ottemann2and Karen Guillemin1
Received 26 February 2011
Revised 28 April 2011
Accepted 12 May 2011
1Institute of Molecular Biology, University of Oregon, Eugene, OR 97403, USA
2Department of Microbiology and Environmental Toxicology, University of California, Santa Cruz, CA
Helicobacter pylori moves in response to environmental chemical cues using a chemotaxis two-
component signal-transduction system. Autoinducer-2 (AI-2) is a quorum-sensing signal
produced by the LuxS protein that accumulates in the bacterial environment in a density-
dependent manner. We showed previously that a H. pylori luxS mutant was defective in motility on
soft agar plates. Here we report that deletion of the luxS gene resulted in swimming behaviour
with a reduced frequency of stops as compared to the wild-type strain. Stopping frequency was
restored to wild-type levels by genetic complementation of the luxS mutation or by addition of
synthetic 4,5-dihydroxy-2,3-pentanedione (DPD), which cyclizes to form AI-2. Synthetic DPD also
increased the frequency of stops in wild-type H. pylori, similar to the behaviour induced by the
known chemorepellent HCl. We found that whereas mutants lacking the chemoreceptor genes
tlpA, tlpC or tlpD responded to an exogenous source of synthetic DPD, the chemoreceptor
mutant tlpB was non-responsive to a gradient or uniform distribution of the chemical. Furthermore,
a double mutant lacking both tlpB and luxS exhibited chemotactic behaviour similar to the tlpB
single mutant, whereas a double mutant lacking both tlpB and the chemotransduction gene cheA
behaved like a nonchemotactic cheA single mutant, supporting the model that tlpB functions in a
signalling pathway downstream of luxS and upstream of cheA. We conclude that H. pylori
perceives LuxS-produced AI-2 as a chemorepellent via the chemoreceptor TlpB.
Directed motility in response to environmental chemical cues,
a process called chemotaxis, is an important trait for many
bacterial pathogens. Helicobacter pylori, a Gram-negative
gastric pathogen, is proposed to situate itself within the gastric
mucosa in response to specific chemical cues such as pH
(Schreiber et al., 2004). Dysregulation of chemotaxis com-
promises the ability of H. pylori to colonize the murine and
gerbil stomach, and results in its abnormal distribution and
modulation of inflammation in this tissue (Foynes et al., 2000;
McGee et al., 2005; Terry et al., 2005; Williams et al., 2007).
The chemotactic response, which has been studied
most extensively in Escherichia coli, is mediated by
proteins (MCPs) (Armitage, 1999). The H. pylori chemo-
sensory machinery is similar but not identical to that of E.
coli. The core signal-transduction apparatus (CheW, CheA
and CheY) is present in both bacteria, but H. pylori also
possesses three CheV proteins composed of both CheW and
response-regulator motifs (Lowenthal et al., 2009; Pittman
et al., 2001). The H. pylori genome encodes four chemor-
eceptors, TlpA, TlpB, TlpC and TlpD (originally called HylB
or HlyB), although strain G27 does not express TlpC due to
a frameshift mutation in the gene. Unlike peritrichous E. coli
cells, which exhibit forward swimming and tumbling
behaviour, we and others have observed that H. pylori cells
swim forward, reverse and stop (Lowenthal et al., 2009;
Schweinitzer et al., 2008). Once the cells stop, they usually
resume movement in a different direction, thus allowing
them to explore new spaces, similar to the tumbling
behaviour of E. coli.
Only a handful of chemotactic signals have been demon-
strated for H. pylori chemoreceptors. In strain 26695 the
chemoreceptor TlpA has been shown to respond to
Abbreviations: AI-2, autoinducer-2;DPD,4,5-dihydroxy-2,3-pentanedione;
iPCR, inverse PCR; SAM, S-adenosylmethionine.
3Present address: Department of Molecular and Cell Biology, University
of Connecticut, Storrs, CT 06269, USA.
Washington, Seattle, WA 98195, USA.
Department of Microbiology,University of
Microbiology (2011), 157, 2445–2455
049353G2011 SGM Printed in Great Britain 2445
arginine and bicarbonate (Cerda et al., 2003), and in strain
SS1 the chemoreceptor TlpB has been demonstrated to
respond to acid (Croxen et al., 2006). The intracellular
chemoreceptor TlpD has been described as an energy
sensor, although the molecular nature of the cues it
perceives is not known (Schweinitzer et al., 2008). There
are no known ligands for TlpC.
One environmental condition that H. pylori experiences is
the endogenously produced quorum-sensing molecule
autoinducer-2 (AI-2) (Forsyth & Cover, 2000; Joyce et al.,
2000; Rader et al., 2007). Autoinducers (AIs) are bacterially
produced extracellular signalling molecules that trigger
quorum sensing, a form of bacterial cell–cell communica-
tion (Ng & Bassler, 2009). AI concentration increases as a
function of bacterial cell density, and at critical concentra-
tion thresholds it initiates coordinated gene regulation.
Several types of quorum-sensing systems have been
characterized. Typically, Gram-positive and Gram-negative
bacteria use oligopeptides and acylated homoserine lactones
as AIs, respectively. Many Gram-positive and Gram-
negative bacteria possess a quorum-sensing system that
produces the furanone signal AI-2, which functions as a
signal for interspecific communication. AI-2 is produced as
a metabolic byproduct of the reaction carried out by LuxS,
which cleaves S-ribosylhomocysteine, producing homocys-
teine and 4,5-dihydroxy-2,3-pentanedione (DPD). DPD
undergoes rapid dehydration and cyclization, existing in
equilibrium as several molecules collectively termed AI-2
(Schauder et al., 2001). In many bacteria, LuxS is a key
enzyme in the production of the activated methyl donor, S-
adenosylmethionine (SAM); however in H. pylori, the
homocysteine produced by LuxS is metabolized to produce
cysteine (Doherty et al., 2010).
Previous reports have shown that several H. pylori strains
produce AI-2 in a luxS-dependent fashion, as detected
through a Vibrio harveyi luminescence assay (Forsyth &
Cover, 2000; Joyce et al., 2000; Lee et al., 2006; Rader et al.,
2007; Shen et al., 2010). Mutation of the luxS gene has
been associated with reduction of the flagellin flaA
transcript and protein, enhanced biofilm formation,
decreased motility on soft agar plates, and colonization
defects in mouse and gerbil infection models (Cole et al.,
2004; Lee et al., 2006; Loh et al., 2004; Osaki et al., 2006;
Rader et al., 2007). We have previously reported that
LuxS-produced AI-2 functions as a signal molecule that
modulates transcript levels of the flagellar regulator FlhA,
thereby influencing global flagellar regulation (Rader et al.,
2007). Although it is clear that AI-2 signalling affects
flagellar gene expression, we found that the flagella in luxS
mutants are morphologically normal and thus the altered
flagellar gene regulation in this strain may not explain
fully its motility defect on soft agar plates. Because
colonial expansion in these plates requires both motility
and chemotaxis, we sought to examine how AI-2 affects
the latter process. In this study we identify a novel
function for AI-2 as a chemorepellent that is sensed by the
Bacterial strains and culture conditions. H. pylori strain G27 and
its isogenic mutants were used in this study and are listed in Table 1.
All H. pylori strains were maintained on blood agar plates consisting
of Columbia agar (Difco) and 5% defibrinated horse blood
(Hemostat) (CHBA), or CHBA plates supplemented with 0.02 mg
b-cyclodextrin ml21(Sigma), 8 mg amphotericin B ml21(Sigma)
and 20 mg vancomycin ml21(Sigma), and incubated at 37 uC in 10%
CO2. Selective plates were supplemented with 10–15 mg kanamycin
ml21(Fisher), 18 mg metronidazole ml21, 10 mg chloramphenicol
ml21or 80 mM sucrose. H. pylori liquid medium (BB10) consisted of
filtered sterilized Brucella broth (Difco) supplemented with 10% fetal
bovine serum (Gibco) and 20 mg vancomycin ml21(Sigma). Liquid
cultures were grown in 50 ml conical tubes (BD Falcon) with
loosened lids shaking at 37 uC in anaerobic jars (Oxoid) with
CampyGen microaerobic sachets (Oxoid) or in glass test tubes
(206148 mm) shaking in a 37 uC/8% CO2incubator.
Table 1. Strains used in this study
H. pylori strainDescription Reference
luxS tlpB (CW30)
tlpB cheA (CW31)
G27, luxS::kan sacB
G27, deletion of luxS
G27, deletion of luxS and rdx::luxS
Mouse-passaged strain G27
mG27, deletion of tlpA
mG27, deletion of tlpB
mG27, deletion of tlpB and rdx::tlpBSS1-aphA-3
mG27. tlpB::cat and luxS::kan sacB
mG27, tlpD::cat (made using pL30A2cat2)
mG27, cheA::cat reverse (made using pKT22)
G27, tlpB::kan sacB and cheA::cat reverse
Covacci et al. (1993)
Rader et al. (2007)
Rader et al. (2007)
Rader et al. (2007)
Castillo et al. (2008)
This study/Williams et al. (2007)
This study/Williams et al. (2007)
This study/Terry et al. (2005)
B. A. Rader and others
Construction of H. pylori mutants. All transformations were
performed using natural transformation. PCR for cloning purposes
was done with either Pfu Turbo (Stratagene) or Phusion (NEB), and
all DNA manipulation enzymes were from New England Biolabs. The
G27 luxS, DluxS, and luxS* isogenic strains (strains BR08, BR09 and
BR10, respectively) were constructed as previously described (Rader et
al., 2007). Throughout, we use ‘D’ to designate deletion mutations, in
which most of a gene’s coding sequence is removed by allelic
exchange, and the superscript ‘*’ to designate complemented strains
in which a wild-type copy of a mutated gene is supplied at another
genomic locus from the mutated gene. The luxS* strain contains a
wild-type copy of the luxS gene and its promoter introduced in the
rdxA locus of the DluxS mutant chromosome. For most chemotaxis
mutants, a mouse-selected variant of G27, mG27, was used as the
parent strain (Castillo et al., 2008).
The cheA mutant (strain KO629) was created by transforming H.
pylori mG27 with pKT22 (Terry et al., 2005) and selecting for
resistance on chloramphenicol CHBA.
DtlpB (strain KO1004) was made in two steps. First, mG27 was
transformed to kanamycin resistance (KmR) using the plasmid pTC-
B111, which bears a tlpB::kan-sacB allele. The correct nature of this
mutation was verified by PCR using primers that flank the insertion
site. pTC-B111 was made starting with pTCB101, which has tlpB from
strain SS1 cloned into pBluescript (McGee et al., 2005). pTCB101 was
then subjected to iPCR using primers tlpB-30 and tlpB-40 (sequence
in McGee et al., 2005). The kan-sacB construct was obtained from
pKSFII (Copass et al., 1997) by digesting with SmaI and XhoI. The
kan-sacB and tlpB flanking pieces were ligated using T4 ligase. The
correct nature of the plasmid was verified by digestion. In this
construct, tlpB is transcribed in the same direction as sacB (opposite
to aphA-3). This strain is called KO1003. KO1003 was then
transformed with a plasmid bearing a large deletion of tlpB, pTC-
B113, followed by selection for sucrose resistance (SucR). pTC-B113
was created using iPCR with primers tlpB-30 and tlpB-40, followed by
intramolecular ligation of the plasmid. The deletion removes all of
tlpB except the first 27 and the last 17 amino acids, although the
deletion is +1 out of frame, resulting in a 39 end that encodes a
different sequence with three additional amino acids. The tlpB region
from SucR, kanamycin-sensitive (KmS) colonies was amplified using
PCR with primers that flank tlpB. This PCR product was sequenced to
verify that tlpB was deleted.
To create the tlpB* complemented strain (strain BR31), mG27 DtlpB
mutant bacteria were naturally transformed with a construct
encoding the rdxA locus with an insertion composed of the full-
length tlpB gene, expressed from the cheY promoter, and cloned
upstream of the non-polar aphA-3 cassette. Disruption of the rdxA
locus confers metronidazole resistance, and the non-polar aphA-3
cassette contains two internal ribosome-binding sites flanking the
kanamycin-resistance gene conferring that resistance. Transformants
were first screened on kanamycin plates, further screened on
metronidazole-supplemented plates, and verified by PCR amplifica-
tion and sequencing of the genomic locus. For the complementation
construct, the non-polar aphA-3 [original sequence from pUC18 K-2
(Me ´nard et al., 1993)] was PCR amplified from the previously
described G27 fliA mutant strain (Rader et al., 2007) using M13F (59-
GTAAAACGACGGCCAGT-39) and M13R (59-CAGAAGACAGCT-
ATGA-39) primers. To create the final DNA construct, a modified
form of the vector pK0140 (Terry et al., 2005) was generated called
pKO140-tlpB in which the cheY coding sequence was replaced by the
tlpB coding sequence from SS1, retaining the cheY promoter
sequence. The aphA-3 DNA and pKO140-tlpB were digested with
SalI and EcoRI and ligated together, creating the rdx:tlpBSS1-aphA-3
construct pK1337. The rdx:tlpBSS1-aphA-3 sequence was then PCR
amplified from pK1337 using the RdxA1F and RdxA2R primers
(Rader et al., 2007). This PCR product was then introduced into the
mG27 DtlpB mutant through natural transformation followed by
selection for KmRisolates. Transformants were further screened on
metronidazole supplemented plates and verified by PCR amplifica-
tion and sequencing of the genomic locus.
The tlpA mutant (strain KO1002) was made in two steps by first
transforming mG27 to KmRwith pTA12, which bears a tlpA::kan-
sacB allele, to create strain KO1001, followed by transformation with
a plasmid that bears a DtlpA deletion (pTA14). pTA12 was made from
pTA10, which has tlpA from strain SS1 cloned, with ~500 bp flanking
sequence, by PCR amplification with primers TlpA6 (59-ATTGAGC-
GCAAAAATAGGGGC-39) and TlpA7 (59 TTTTCTCTCGCCAAAG-
CTTGC-39). This piece was phosphorylated with T4 polynucleotide
kinase, and then cloned into EcoRV-digested pBluescript KS+
(Stratagene) to create pTA10. To create pTA12, iPCR was carried
out with pTA10 template plus primers TlpA12 (59-CACCCACAATA-
TGATTTTATTACCG-39) and TlpA13 (59-CAAGAAATTGACAAA-
GTCTCTAACG-39) to create an in-frame deletion that leaves a region
coding for 14 amino acids at the 59 end, and 19 amino acids at the 39
end of tlpA. This iPCR product was then ligated to a kan-sacB
fragment that came from pKS2. pKS2 derived from pKS1, which was
made via a three-piece ligation of the following pieces: (1) the aphA3
gene (also called kan) isolated from pBS-kan (Terry et al., 2005) by
cutting with SmaI+XhoI, followed by dephosphorylation; (2) the
sacB gene from pKSFII (Copass et al., 1997) generated by cutting with
BamHI, blunting with T4 polymerase followed by cutting with PstI;
(3) the vector backbone from pBluescript KS+ cut with PstI and
XhoI, followed by phosphatasing and gel purification of the 3.0 kb
piece. These three pieces were ligated together to generate pKS1. pKS1
has the aphA3 promoter driving transcription of both aphA3 and the
downstream sacB gene. To create pKS2, pKS1 was digested with PvuI
to remove the bla gene followed by self-ligation. For cloning of the
aphA3-sacB fragment from pKS2, PCR amplification using primers
reverse (universal) and sacBend2 (59-CTTTTGCGTTTTTATTTG-
TTAAC-39) was carried out, to amplify aphA3-sacB without the sacB
transcriptional terminator, and the resulting fragment was ligated
with the iPCR fragment of pTA10 to create pTA12. In pTA12, tlpA,
aphA3 and sacB are all in the same transcriptional orientation. To
create pTA14, the same pTA10 iPCR fragment was self-ligated. H.
pylori KO1001 was then transformed with pTA14, followed by
selection for sucrose resistance and screening for kanamycin
sensitivity. The tlpA allele was PCR amplified from SucRKmS
colonies using primers tlpA10 (59-TCTAAAGGTTTGAGTATCGG-
39) and tlpA11 (59-GCTCGAATTCGAAAACTGCTTTTTATTCAC-
ATC-39), and sequenced to verify the correct deletion. This tlpA strain
is also called mG27 tlpA or KO1002.
The mG27 tlpC mutant is tlpC::aphA3 (strain KO1005). This mutant
was constructed by transforming mG27 with plasmid pKO150. This
plasmid was made using iPCR of a plasmid containing tlpC from SS1
called pTC100 (Andermann et al., 2002) with primers tlpC31 (59-
TCATCACAATTTTAGAACC-39) and tlpC40 (59-CCTTGCAACAA-
GATGTGCAGG-39). The product, 3.5 kb, was ligated with the aphA3
gene prepared by PCR from pBS-kan with primers Kanup (59-
GGCCGGATCCGATAAACCCAGCGAACC-39) and Kandown (59-
GGCCAAGCTTTTTAGACATCTAAATC-39). The resulting product
was phosphorylated with T4 kinase, and ligated to the pTC100DtlpC
product to create pKO150. The tlpD mutant was made by
transforming mG27 to chloramphenicol resistance using plasmid
pL30A2cat2 (Williams et al., 2007), which creates tlpD::cat. Note
that tlpD was previously referred to as hylB.
To generate strain CW30, with mutations in both luxS and tlpB genes,
we transformed H. pylori mG27 tlpB::catR (Williams et al., 2007)
with PCR DNA generated from luxS::kan sacB genomic DNA and
selected for kanamycin resistence. This luxS::kan-sacB DNA was
amplified with primers luxSfor (59-AACGCTGGGATTACGCATG-
GA-39) and luxSrev (59-AAGCCGCCCGTGAATGTCTGAA-39).
Autoinducer-2 is a chemorepellent for H. pylori
To generate the bacterial strain CW31 with mutations in both cheA
and tlpB genes, we first created a tlpB::kan-sacB mutation in wild-
type G27 bacteria and then added the cheA::cat reverse mutation. To
generate the DNA for the tlpB::kan-sacB mutation, SS1 tlpB was
amplified from the pK1337 plasmid using SS1TlpBF (59-TAAG-
GCGTTAGAGACGCTTTGGCT-39) and SS1TlpBR (59-AAACACG-
CCGTGATCACAGAAACC-39) primers and ligated into the pCR 2.1-
TOPO plasmid using the TOPO TA Cloning kit (Invitrogen), creating
pTOPO-tlpB. The kan-sacB cassette (Copass et al., 1997) was
amplified from the pKSF-II plasmid using primers kansacBF (59-
CTCCATGGTCCCGGGCGAACCATTTGAGGTGA-39) and kansacbR
The kan-sacB cassette was inserted into the HincII restriction site at
tlpB::kansacB. The tlpB::kan-sacB sequence was amplified from
pTOPO-tlpB::kansacB and transformed into naturally competent G27
H. pylori. Positive tlpB::kan-sacB transformants were selected by
kanamycin resistance and confirmed via PCR using primers TlpBF (59-
ACTTCAAAAGACGGGAGGACT-39) and TlpBR (59-TATCCCCAC-
TCGCACGC-39). The tlpB::kan-sacB bacteria were transformed with
PCR DNA generated from cheA::cat reverse genomic DNA, and
selection for chloramphenicol resistance was applied. The correct
genotype was confirmed by PCR. The cheA::cat reverse DNA, was
amplified with primers CheA_for1 (59-GTGCTGAAAGGGCTA-
AAGAAATG-39) and CheA_rev1 (59-GGATAATCGCTCTGTCCG-
Bioluminescence assay and synthetic DPD. Synthetic DPD, a
kind gift from Martin Semmelhack and Bonnie Bassler (Princeton
University), was prepared as described by Semmelhack et al. (2005).
This DPD was synthesized with a protecting group that we removed
by treatment with 15 mM H2SO4, followed by neutralization to
pH 6.9 with potassium phosphate. As a mock DPD treatment, we
neutralized 15 mM H2SO4 to pH 6.9 with potassium phosphate.
Alternatively, we used DPD from OMM Scientific that did not
contain a protecting group; BB10 medium was used to dilute this
DPD and as a mock treatment. The concentration of biolumin-
escence-inducing activity in H. pylori cell-free supernatant from early
stationary-phase cultures was approximately equivalent to 0.1 mM
synthetic DPD (Rader et al., 2007).
Microscopy. Swimming behaviour of bacterial strains was visualized
using an inverted Leica DMIL phase-contrast light microscope (Leica
Microsystems). G27 wild-type, DluxS, luxS*, tlpB and tlpB* isogenic
strains were grown overnight on a shaker to an OD600of 1.0. Aliquots
(2 ml) of this overnight culture were then added to a test tube
containing 500 ml fresh BB10 medium and placed stationary at 37 uC
in 10% CO2 for 2 h. These conditions promoted high levels of
motility. Approximately 8 ml of the stationary culture was spotted
onto a glass slide followed by the same amount of 0.1 M HCl, 0.1 mM
synthetic DPD, mock DPD solution, or BB10 medium as a negative
control. A coverslip was added and sealed on all sides with clear
finger-nail polish. After 4–10 min, videos were recorded at 15 or 30
frames s21on a Cohu High Performance CCD camera using Scion
Image software. The number of stops exhibited by individual bacteria
was determined by examining 5 s of swimming for multiple bacterial
cells per video, as described previously (Terry et al., 2006). Stops were
defined as the cessation of movement in any direction for three or
more frames; we did not score reversals that occurred without pauses.
Individual bacterial cells were scored if they met the following criteria:
they were motile at the beginning of the 5 s interval, they remained
visible for the duration of the 5 s interval, and they stopped only for
at most 30 frames, and resumed swimming after this period. Video
scoring was performed without knowledge of the genotype or
treatment condition. At least two independent trials were performed
per condition and at least 15 bacterial cells were scored per condition.
To assay bacterial taxis away from a source of chemical repellent, we
adopted the previously described barrier formation assay (Croxen
et al., 2006). Wild-type mG27, DtlpA, DtlpB, tlpB*, tlpC, tlpD and
cheA isogenic strains were grown overnight, with shaking, in BB10 to
an OD600of 1.0, followed by 2 h without shaking, as above. Samples
(8 ml) of culture were spotted onto the centre of a glass slide and
covered with a no. 1, 22622 mm wide glass coverslip. Three sides of
the coverslip were sealed with clear finger-nail polish. Eight
microlitres of 0.1 M HCl, 0.1 mM synthetic DPD, or mock DPD
solution was placed on the open side of the coverslip. After 5–10 min
incubation, the slides were examined and still images were taken using
Scion Image software at 610 through a 640 phase filter, conditions
that mimic dark-field microscopy.
Western blotting. Total cell proteins were prepared from H. pylori
cultured on CHBA plates for 2 days by resuspending and lysing the
cells in 26 Laemmli sample buffer. Samples were separated on a 10%
SDS-PAGE gel, transferred to Immunoblot PVDF membranes (Bio-
Rad) and incubated with 1:5000 dilution of anti-GST_TlpA22
(Williams et al., 2007). This rabbit polyclonal antibody recognizes the
conserved CheW-interacting methyl-accepting domain of all H. pylori
chemoreceptors. For visualization, the blots were incubated with the
secondary antibody goat anti-rabbit-HRP (Santa Cruz Biotech) at a
dilution of 1:2000, followed by incubation with luminol, p-coumaric
acid and hydrogen peroxide. Luminescent blots were visualized by
exposure to Biomax Light film (Kodak).
Statistical analysis. The frequency of stops was analysed statistically
using either one-way ANOVA (www.physics.csbsju.edu/cgi-bin/stats/
anova_pnp) when strains were tested or two-way ANOVA (http://
faculty.vassar.edu/lowry/anova2u.htl) when both strains and chemical
treatment were tested. Post-hoc multiple comparisons to identify
significance within individual hypotheses were performed using t-
tests, and the P-values were adjusted using Bonferroni correction to
correct for an increase in type I error. P-values of ,0.001 were
considered significant and used for Bonferroni correction.
luxS is required for normal swimming behaviour
of H. pylori
When examining the swimming behaviour of wild-type
and an isogenic luxS deletion mutant (referred to as DluxS)
of H. pylori strain G27 (Rader et al., 2007), we observed
DluxS mutant displayed a propensity for
swimming in a straight line (running), as opposed to the
wild-type, which exhibited more typical run–stop–run
behaviour. We quantified this behaviour in bacterial
cultures using video microscopy to observe swimming
behaviour. We found that the DluxS mutant exhibited
significantly fewer stops per second than the wild-type
strain (Fig. 1). Complementation of the luxS mutation with
a wild-type copy of the luxS gene at the rdxA locus [referred
to as luxS* (Rader et al., 2007)] significantly increased the
number of stops per second as compared with the DluxS
strain (Fig. 1). It should be noted that over the course of
these experiments, we observed some variation between
trials in the average number of stops per second exhibited
by the strains, but the responses to chemicals and
differences between genotypes were reproducible across
experiments. We attribute these differences to factors that
B. A. Rader and others
were difficult to control across all experiments, such as
differences in the lots of serum used in the media.
The LuxS product, AI-2, is a chemorepellent for H.
To test whether the decrease in the number of stops in the
DluxS mutant relative to the G27 wild-type and luxS*
isogenic strains was due to the loss of the AI-2 signal, we
treated each strain with 0.1 mM synthetic DPD, corres-
ponding to the approximate concentration of AI-2 in the
cell-free supernatant of a early stationary-phase culture of
wild-type H. pylori (Rader et al., 2007). We also treated
each isogenic strain with sterile water, the DPD diluent, as
a negative control. Bacterial cultures were incubated with
treatment for 10 min before video microscopy. With the
addition of synthetic DPD, the number of stops in the DluxS
mutant was increased to a level similar to that seen in the
wild-type strain treated with water (Fig. 1). Interestingly, in
the cultures of the wild-type and the complemented strains,
the number of stops also increased in the presence of
synthetic DPD, suggesting that all three isogenic strains
responded to changes in AI-2 concentration.
The swimming behaviours exhibited in all three isogenic
strains upon addition of synthetic DPD were strikingly
similar to the reported repellent response of H. pylori strain
SS1 to the chemorepellent HCl (Croxen et al., 2006). To
investigate whether strain G27 responded similarly to HCl
and synthetic DPD, we incubated wild-type, DluxS and
luxS* G27 with 0.1 M HCl and observed their swimming
behaviour. The pH of the synthetic DPD solution was 6.9,
in contrast to the pH of 1.0 of the 0.1 M HCl solutions. All
three isogenic strains exhibited an increase in the number
of stops when exposed to 0.1 M HCl, and this increase did
not differ significantly from that in response to synthetic
DPD (Fig. 1). To ensure that this similarity was not due to
some component of the solution used to rehydrate and
activate the synthetic DPD, we also performed the
experiment with a solution identical to the synthetic
DPD solution, to which the synthetic DPD was not added
(mock DPD). This treatment did not alter the swimming
behaviour of the strains relative to their behaviour in
response to water (Fig. 1). All subsequent experiments
employed mock DPD as the negative control. These
experiments thus suggest that DPD functions as a
chemorepellent for H. pylori, because it increases the
frequency of bacterial stopping.
To further explore whether H. pylori uses chemotaxis to
respond to synthetic DPD, we employed a previously
described wet-mount assay to monitor spatial chemotactic
responses (Croxen et al., 2006). Wild-type G27 bacterial
suspensions in BB10 medium were prepared as described
earlier, seeded onto a glass microscope slide, and covered
with a coverslip that was then sealed on three sides.
Synthetic DPD (0.1 mM), HCl (0.1 M), mock DPD or
water was then spotted onto the part of the coverslip that
remained unsealed (Fig. 2a). Movement of the bacterial
population was monitored before, immediately after and
5 min after addition of the chemical stimulus. Prior to the
addition of treatment the bacteria were motile and
uniformly distributed below the coverslip. Immediately
after addition of treatment, the bacteria were flushed away
from the liquid source; however after 5 min the hydro-
dynamic flow had ceased, and in the case of mock DPD or
water, the bacteria returned to a uniform distribution. As
previously reported (Croxen et al., 2006), in response to
0.1 M HCl, the percentage of motile bacteria immediately
increased, with the majority of the bacteria exhibiting an
increase in the frequency of stops, resulting in net
movement away from the source of the acid. This increased
movement proximal to the acid source ended in the
formation of a visible barrier of bacterial cells toward the
centre of the coverslip that accumulated presumably at a
preferred concentration of the chemical treatment (Fig.
2C). Beyond this barrier, bacterial movement resembled
that of the untreated controls. These observations show
that H. pylori strain G27 responds to pH similarly to strain
SS1 (Croxen et al., 2006). When treated with 0.1 mM
synthetic DPD, the cultures exhibited a response that was
similar to that to acid: the cells showed increased frequency
of stops and formed a bacterial barrier at a similar location
away from the chemical source (Fig. 2D). In response to
mock DPD, the percentage of motile bacteria in the
population increased, but there was no obvious net
directional movement, and after 5 min no bacterial barrier
Fig. 1. LuxS is required for normal swimming behaviour of H.
pylori. Swimming behaviours of G27 wild-type, DluxS mutant and
luxS* complemented isogenic strains were observed by video
microscopy, and the number of stops that individual bacteria
performed during 5 s was recorded. Each strain was treated with
water, mock DPD, 0.1 M HCl and 0.1 mM synthetic DPD. The
number of individual bacterial cells scored (n) for each condition is
indicated below the column; bars indicate SE. * indicates each
strain that is statistically significant different from all other strains
within a treatment, + indicates chemorepellent treatments (HCl,
synthetic DPD) that are significantly different from controls (water,
mock DPD) within a strain (P,0.001 with Bonferroni correction).
Autoinducer-2 is a chemorepellent for H. pylori
had formed and the bacteria were evenly distributed
underneath the coverslip (Fig. 2B). Our data thus suggest
that H. pylori cells respond chemotactically to synthetic
DPD similarly to how they respond to HCl.
The chemotaxis receptor TlpB is required for
chemotaxis away from AI-2
Bacterial tactic movement toward or away from a chemical
source is generallyregulated bythe chemotaxis system. Key to
this system are the chemoreceptors which sense envi-
ronmental signals and transmit the information through a
transduction cascade, eventually affecting flagellar rotation
(Armitage, 1999). To test whether the response to synthetic
DPD was due to detection by any of the four H. pylori
chemoreceptors, we repeated the wet-mount assay described
above, using mouse-passaged H. pylori mG27 isogenic strains
harbouring deletions or insertional mutations in the
chemoreceptor genes tlpA (strain DtlpA), tlpB (strain
DtlpB), tlpC (strain tlpC) or tlpD (strain tlpD). The wild-
type strain mG27 does not produce TlpC as detectable by
Western blotting (Fig. 3A), but we included the engineered
tlpC mutant in our analysis for completeness. Each of these
strains was challenged with 0.1 mM synthetic DPD, 0.1 M
HCl or mock DPD. As a negative control for chemotaxis we
employed a non-chemotactic mutant with a mutation in the
chemotransduction gene cheA. The DtlpA, tlpC and tlpD
mutants all formed bacterial barriers when challenged with
reported, the DtlpB mutant failed to form a bacterial barrier
when challenged with HCl (Fig. 2I). In addition, this mutant
failed to form a bacterial barrier when challenged with
syntheticDPD(Fig.2J).Asexpected,the cheAmutant didnot
form a bacterial barrier in response to HCl or synthetic DPD
(Fig. 2R, S). None of the five strains produced bacterial
barriers in response to mock DPD (Fig. 2E, H, K, N, Q).
These data implicated TlpB in the sensing of AI-2.
To verify that the failure of the DtlpB mutant to form a
bacterial barrier in response to DPD was due to the lack of
the tlpB gene product, we constructed a complemented
strain (referred to as tlpB*) by placing the full-length tlpB
gene in the rdxA locus of the DtlpB mutant chromosome.
Western blot analysis verified production of the TlpB
protein in the tlpB* strain (Fig. 3A). We then repeated the
wet-mount assay with the tlpB* strain. When challenged
with 0.1 mM synthetic DPD or 0.1 M HCl, the tlpB* strain
responded by producing a bacterial barrier, in contrast to
the mock DPD solution, which produced a uniform
distribution of bacteria (Fig. 3B).
To further characterize TlpB as a chemoreceptor for
synthetic DPD, we repeated the video taxis assay with the
mG27 wild-type, DtlpB and tlpB* isogenic strains. The
average number of stops for each genotype was recorded
for each strain grown in mock DPD, 0.1 M HCl or 0.1 mM
synthetic DPD. As reported previously (Croxen et al.,
2006), we observed that the DtlpB mutant displayed an
increased stopping frequency as compared to its wild-type
parent in medium without added chemorepellents (see
below). To best represent the relative responsiveness of the
different strains to HCl and DPD, we therefore show the
normalized stopping frequencies for each strain in each
condition relative to the average number of stops exhibited
by that strain in the mock DPD medium (Fig. 3C). In
accordance with the wet-mount assay, the DtlpB mutant
exhibited significantly reduced responsiveness to DPD and
HCl as compared with the wild-type strain. The comple-
mented tlpB* strain had restored responsiveness to DPD
Fig. 2. Chemotaxis response of wild-type and mutant H. pylori to
HCl and synthetic DPD. A wet-mount chemotaxis assay (illustrated
in A) was used to analyse chemotatic behaviour of mG27 wild-type
and the indicated isogenic mutant strains in response to mock
DPD, the known chemorepellent 0.1 M HCl, and 0.1 mM synthetic
DPD. White arrows indicate bacterial barrier formation.
B. A. Rader and others
2450 Microbiology 157
and greater responsiveness to acid than the wild-type
strain. We do not know the reason for this strain’s hyper-
responsiveness to acid, but it could be due to altered
expression of the tlpB gene under acidic conditions when
expressed from a different promoter and genomic locus
than those of the native gene.
As noted above, the DtlpB mutant displayed an increased
stopping frequency as compared to its wild-type parent
when grown in medium in the absence of exogenously
increased frequency was due to a hyper-activation of the
chemotaxis signal transduction pathway, or some other
effect on swimming behaviour caused by the loss of tlpB. To
distinguish these possibilities, we constructed a tlpB cheA
double mutant and compared the swimming behaviour of
this strain to the tlpB and cheA single mutants. As expected,
the cheA single mutant, which lacks a functional chemotaxis
signal tranduction pathway, exhibited fewer stops than the
wild-type strain in BB10 (Fig. 4A). The tlpB cheA double
mutant was indistinguishable from the cheA single mutant,
indicatingthat the increased stopping observedwith the tlpB
mutant was due to increased activation of the chemotaxis
pathway, possibly as a result of increased signalling from
other chemoreceptors in the absence of TlpB.
TlpB functions downstream of LuxS in AI-2
Our data suggest that TlpB functions as the chemorecep-
tor for avoidance responses to AI-2, a molecule produced
by the bacteria themselves using the enzyme encoded by
luxS. If this model were correct, then we would predict
that cells lacking both LuxS, the enzyme that makes AI-2,
and TlpB, the chemoreceptor involved in its perception,
should resemble cells lacking only the receptor. We
generated a luxS tlpB double mutant and compared its
swimming behaviour to that of luxS and tlpB single
mutants. We observed that the luxS tlpB double mutant
exhibited a high stopping frequency, statistically indistin-
guishable from the tlpB single mutant (Fig. 4A).
Importantly, whereas the luxS single mutant formed
barriers to both DPD and HCl in the wet-mount assay
(Fig. 4F, G), the luxS tlpB double mutant failed to form
any barriers (Fig. 4I, J), similar to the tlpB single mutant
Fig. 3. TlpB is required for chemotactic responses to HCl and synthetic DPD. (A) Western blot analysis of mG27 wild-type,
DtlpB, and tlpB* isogenic strains with an antibody that recognizes a conserved region of all H. pylori chemoreceptors. (B)
Chemotactic behaviour of the tlpB* complemented strain was assayed using the wet-mount assay in response to mock DPD,
0.1 M HCl and 0.1 mM synthetic DPD. (C) The normalized stopping frequency for mG27 wild-type, DtlpB and tlpB* isogenic
strains treated with mock DPD, 0.1 M HCl and 0.1 mM synthetic DPD is shown, where the average number of stops for each
strain in each condition is normalized to that strain’s average number of stops in mock DPD. The number of individual bacterial
cells scored (n) for each condition is indicated below the column; bars indicate SE. * indicates statistically significant differences
in stopping frequency as compared to the mock DPD treatment of the same strain (P,0.001 with Bonferroni correction).
Autoinducer-2 is a chemorepellent for H. pylori
(Fig. 2I, J). These data demonstrate that tlpB is epistatic to
luxS, consistent with our model that TlpB perceives a
molecule produced by the luxS gene product.
In this study we have shown that the quorum-sensing
signal AI-2 acts as a chemorepellent for H. pylori, and
demonstrated that the chemoreceptor TlpB is required for
its perception. We previously reported that elimination of
the luxS gene product in H. pylori exhibited a motility
defect on soft agar (Rader et al., 2007), which was
complemented by exogenous AI-2, a result confirmed by
others (Shen et al., 2010). This assay cannot distinguish
between dysfunction of the motility apparatus and
dysregulation of chemotaxis. To better characterize the
chemotactic response of H. pylori to the absence or
presence of AI-2, we observed both swimming behaviour
of individual bacterial cells and the response of populations
of bacteria to chemical gradients. We found that luxS-
deficient bacterial cells exhibited a decrease in stopping
frequency as compared to wild-type cells. This behaviour
was restored to wild-type levels by the addition of synthetic
DPD. Synthetic DPD also increased the frequency of stops
in the wild-type strain, and caused populations of wild-
type cells to move away from a source of the chemical,
suggesting that AI-2 is a chemorepellent for H. pylori. Our
ability to modulate H. pylori swimming behaviour with
exogenous AI-2 argues against a metabolic requirement for
the luxS gene in chemotaxis. Additionally, although many
bacteria regulate the sensitivity of their chemoreceptors via
methylation, a mechanism which could be influenced by
function of the SAM pathway, H. pylori lacks the methyl-
utilizing enzymes required for this process (Szurmant &
Ordal, 2004). We further demonstrated that an H. pylori
strain deficient for the chemoreceptor TlpB failed to move
away from a source of synthetic DPD, and did not display
increased stopping behaviour upon addition of synthetic
DPD. These behaviours were restored upon genetic
complementation of the tlpB gene.
TlpB and AI-2 perception
Most reported responses to AIs involve regulation of
transcription. Recently, however, E. coli was shown to
respond to AI-2 as a chemoattractant through a mech-
anism that involves both the AI-2 periplasmic binding
protein LsrB and the chemoreceptor Tsr (Bansal et al.,
2008; Hegde et al., 2011). Here we show that H. pylori also
perceives AI-2 as a chemical cue in response to which it
directs its movement, in this case by moving away from the
signal. We demonstrate that the H. pylori chemoreceptor
TlpB is required for perception of AI-2 and confirm its role
in negative pH taxis. Bacterial chemoreceptors can often
sense more than one ligand and can elicit both positive and
negative behavioural responses. For example, the E. coli
chemoreceptor Tsr not only senses AI-2, but also mediates
taxis toward the attractants serine and related amino acids,
and taxis away from weak acids, indole and leucine, and
transduces oxygen and redox signals (Boyd & Simon, 1982;
Rebbapragada et al., 1997). We do not yet know whether
TlpB senses AI-2 directly or via other binding proteins.
TlpB shares no sequence similarity with the previously
identified AI-2 binding proteins, V. harveyi LuxP (Chen
et al., 2002), S. typhimurium LsrB (Miller et al., 2004) and
Aggregatibacter actinomycetemcomitans RbsB and LsrB
(Shao et al., 2007), and the H. pylori genome lacks
homologues of any of these genes. We believe that the
molecular mechanism of chemotaxis from AI-2 is distinct
Fig. 4. tlpB functions in a signalling pathway downstream of luxS
and upstream of cheA. (A) Swimming behaviours in BB10 of G27
wild-type, tlpB, cheA, tlpB cheA, luxS and luxS tlpB isogenic
strains were observed by video microscopy, and the number of
stops that individual bacteria performed during 5 s was recorded.
The number of individual bacterial cells scored (n) for each strain is
indicated below the column; bars indicate SE. Symbols (*, +)
indicate strains that are statistically indistinguishable from each
other (P.0.001 with Bonferroni correction). (B–J) The wet-mount
chemotaxis assay was used to analyse chemotatic behaviour of the
indicated isogenic mutant strains in response to mock DPD, the
known chemorepellent 0.1 M HCl and 0.1 mM synthetic DPD.
White arrows indicate bacterial barrier formation.
B. A. Rader and others
2452 Microbiology 157
from AI-2-mediated regulation of flhA transcription in H.
pylori, because the latter does not require tlpB (B.A.R. and
K.G., unpublished results). In addition, neither tlpB nor
several other chemotaxis genes appear to be regulated
transcriptionally by AI-2 because their transcript levels are
the same in wild-type and DluxS mutant strains (Rader
et al., 2007; B.A.R. and K.G., unpublished results).
AI-2-regulated motility is likely to be important in
the gastric environment
For H. pylori, motility is essential for colonization of piglet,
gerbil and mouse stomachs, where bacterial interactions
with the mucosa promote gastric pathology (Eaton et al.,
1992; McGee et al., 2002; O’Toole et al., 2000; Ottemann &
Lowenthal, 2002). It is well established that non-chemo-
tactic mutants (Che2) of H. pylori do not infect mice to the
full wild-type level (Foynes et al., 2000; Terry et al., 2005).
These mutants, lacking cheA, cheW or cheY, all engage
almost exclusively in straight runs without stops or
changes in direction. We found that the luxS mutant
exhibits a similar swimming behaviour to the Che2
mutants in broth, suggesting that LuxS is responsible for
production of a significant proportion of chemorepulsive
signals present in H. pylori batch culture. We have
confirmed previous reports that a luxS mutant is defective
in colonization of the rodent stomach; however in these
experiments it was not possible to determine whether the
colonization defect was due to reduced AI-2 concentra-
tions in the mouse stomach or metabolic deficiencies in
the bacteria (Lee et al., 2006; Osaki et al., 2006), (B.A.R.,
K.G. and K.M.O., unpublished results). Whereas the
DtlpB mutant colonizes wild-type mice and gerbils to
normal levels (McGee et al., 2005; Williams et al., 2007), it
is impaired in its ability to colonize the stomachs of IL-12-
deficient mice (Croxen et al., 2006). There are likely to be
additional chemical cues to which H. pylori responds
within a mouse stomach (Schreiber et al., 2004). Indeed,
non-chemotactic mutants are less closely associated with
mouse gastric epithelia and induce a diminished inflam-
matory response as compared to wild-type H. pylori,
suggesting that chemical cues from the epithelium are
important for directing H. pylori localization within the
stomach (Williams et al., 2007).
The fact that gastrointestinal pathogens have chemotactic
responses to AI-2 raises the question of the role of this
response in host colonization. In the case of enteropatho-
genic E. coli, the pathogen perceives AI-2 as an attractant,
possibly using it as a cue to direct itself toward the bacteria-
dense colon. We hypothesize that H. pylori perceives AI-2
both as a repellent and as a cue to regulate flagellar gene
expression as a means to coordinate its distribution and
motility within the stomach. We imagine that at some
threshold level of AI-2, the bacteria respond to this
molecule as a chemorepellent and move away from the
bulk bacterial population, thereby avoiding niche com-
petition and promoting dispersal throughout the stomach.
Consistent with this model, luxS mutants in several H.
pylori strains have been reported to form biofilms more
readily than their wild-type counterparts (Cole et al.,
2004). With sustained AI-2 levels, indicative of high
bacterial populations, the elevated AI-2 may alter FlhA
levels to downregulate motility. Such a response might
allow bacteria to avoid wasting energy on swimming and
instead focus on adherence. Several bacterial species
perform reciprocal regulation of motility and adherence
(Holden & Gally, 2004), although this type of response has
not been shown for H. pylori. Furthermore, our under-
standing of the gastric environment has changed in part
due to 16S rRNA gene enumeration studies, which
identified over 120 phylotypes of bacteria in the human
stomach, many of which potentially produce AI-2 (Bik
et al., 2006; Federle & Bassler, 2003). It is possible that H.
pylori will move away from AI-2 produced by coincident
bacteria, providing a further means of avoiding niche
competition. As disease outcome is correlated with
localization of H. pylori populations within the stomach
(Blaser & Atherton, 2004), AI-2 may be an important
environmental factor in the progression of disease in H.
pylori infections. Further understanding of those factors
that regulate H. pylori motility and chemotaxis within the
gastric environment will undoubtedly enhance our ability
to predict disease outcome and design therapies that can
control H. pylori-caused chronic inflammation and sub-
sequent progression to gastric cancer.
This work was supported by National Institutes of Health Public
Health Service grants R01 AI050000 (to K.M.O.) and R01 DK075667
(to K.G.). We thank Bonnie Bassler and Martin Semmelhack for the
generous gift of synthetic DPD, Tessa Andermann and Yu-Ting Chen
for construction of the tlpA and tlpC mutational vectors, and Will
Finch for construction of the pKO140-tlpB vector. We thank
Khoosheh Gosnik, Jim Remington and members of the Guillemin
lab for fruitful discussions.
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Edited by: M. F. Hynes
Autoinducer-2 is a chemorepellent for H. pylori