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Journal of Bacteriology, December 2001, p. 7213-7223, Vol. 183, No. 24
Unité de Microbiologie et
Génétique (CNRS UMR 5122), Institut National des Sciences
Appliquées de Lyon, 69621 Villeurbanne Cedex,
France,1 and Swiss Federal Institute of
Environmental Technology (EAWAG), CH-8600 Dübendorf,
Switzerland2
Received 18 May 2001/Accepted 19 September 2001
The Escherichia coli OmpR/EnvZ two-component
regulatory system, which senses environmental osmolarity, also
regulates biofilm formation. Up mutations in the ompR
gene, such as the ompR234 mutation, stimulate laboratory
strains of E. coli to grow as a biofilm community rather
than in a planktonic state. In this report, we show that the OmpR234
protein promotes biofilm formation by binding the csgD
promoter region and stimulating its transcription. The
csgD gene encodes the transcription regulator CsgD,
which in turn activates transcription of the csgBA
operon encoding curli, extracellular structures involved in bacterial
adhesion. Consistent with the role of the ompR gene as
part of an osmolarity-sensing regulatory system, we also show that the
formation of biofilm by E. coli is inhibited by
increasing osmolarity in the growth medium. The ompR234
mutation counteracts adhesion inhibition by high medium osmolarity; we
provide evidence that the ompR234 mutation promotes
biofilm formation by strongly increasing the initial adhesion of
bacteria to an abiotic surface. This increase in initial adhesion is
stationary phase dependent, but it is negatively regulated by the
stationary-phase-specific sigma factor RpoS. We propose that this
negative regulation takes place via rpoS-dependent
transcription of the transcription regulator cpxR;
cpxR-mediated repression of csgB and
csgD promoters is also triggered by osmolarity and by
curli overproduction, in a feedback regulation loop.
In natural
environments, bacteria are often found as sessile communities known as
biofilms. Biofilms are defined as matrix-enclosed communities of
microorganisms tightly interacting with each other and, in most cases,
supported by an abiotic surface (7). Biofilms can become
hundreds of micrometers in depth and display complex structural and
functional architecture (8, 28, 40). Bacteria growing as a
biofilm develop significant phenotypical, biochemical, and
morphological differences from their planktonic counterparts. In
particular, cells growing in biofilm are extremely resistant to
treatment with biocides and to prolonged antibiotic therapy in human
infections (16, 23). This different biochemical and phenotypical behavior reflects different patterns of gene expression compared with planktonic cells (39, 41). Such
reprogramming of gene expression is likely to be due to changes in
environmental physicochemical conditions and to involve two-component
systems such as EnvZ/OmpR (41) and CpxA/CpxR
(15).
In previous reports we described the isolation of a mutant from a
continuous culture of Escherichia coli K-12 that is more efficient in biofilm formation. Using both genetic analysis and electron microscopy observations, we showed that this increased efficiency was due to augmented production of curli (40,
50). The genes necessary for curli production are clustered in
the csgBA and csgDEFG operons, which encode the
curli subunits and regulate their transcription and transport,
respectively. The csgBA and csgDEFG operons
appear to be expressed in environmental and clinical isolates of
E. coli, as well as in Salmonella strains, in
which the homologous genes are called agfBA and
agfDEFG.
However, some E. coli K-12 laboratory strains do not express
curli, although functional copies of the genes are still present (5, 19, 35). The reason for this different behavior in E. coli K-12 strains is not yet fully understood. Curli
fibers are highly conserved between Salmonella species as
well as in E. coli with respect to curlin amino acid
sequence, genetic organization, and operon regulation (6,
44). Curlin, the product of csgA, is the major
component of curli, while CsgB acts as a nucleator which primes the
polymerization of curlin on the cell surface (20). The
first gene of the csgDEFG operon encodes the CsgD protein, a
putative transcription factor belonging to the luxR family
and required for the transcription of csgBA
(19). The csgEFG genes encode three curlin
assembly factors, probably involved in export of the curli subunits
(19).
Curli biogenesis is subject to tight and complex regulation: in
E. coli K-12 and in Salmonella enterica serovar
Typhimurium, they are only produced at temperatures below 30°C, at
low osmolarity, and in stationary phase (19, 30, 35). The
stationary-phase-induced transcription of genes required for curli
biogenesis is dependent on the In this report we show that curli biosynthesis is subject to a complex
regulatory network: the OmpR protein positively regulates curli
expression by binding the csgDEFG promoter region at
position Bacterial strains, plasmids, and media.
The E. coli strains and plasmids used in this work are listed in Table
1. Media used were Luria-Bertani broth
(LB), minimal M63 medium supplemented with glucose (0.2%)
(32), and M63/2, a low-osmolarity medium obtained by
twofold dilution of the M63 medium and supplementation with glucose
(0.2%). Congo red indicator plates were prepared as described by
Hammar et al. (19); on these plates, curli-producing
bacteria form red colonies, whereas non-curli-producing cells produce
white colonies.
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.24.7213-7223.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
Complex Regulatory Network Controls Initial Adhesion and Biofilm
Formation in Escherichia coli via Regulation of the
csgD Gene
ABSTRACT
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
INTRODUCTION
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
factor RpoS (35).
However, in the absence of the H-NS histone-like protein, transcription
from the csgDEFG promoter becomes independent of RpoS
(3), suggesting that H-NS might selectively repress
70-dependent transcription of csgD.
The two-component systems OmpR/EnvZ (44, 50) and CpxA/CpxR
are also implicated in the regulation of curli biogenesis
(15). The OmpR/EnvZ system constitutes a signal
transduction pathway that senses external osmolarity and regulates the
transcription of several genes, including the porin-encoding genes
ompF and ompC (38). There is genetic
evidence that curli-encoding genes are members of the OmpR regulon
(44, 50), but the existence of direct transcriptional
control has never been demonstrated.
49.5 relative to the transcriptional start site and by
activating its transcription. However, csgDEFG and
csgBA expression is also subject to negative regulation by
the rpoS gene. Negative control by rpoS appears
to be mediated by direct interaction between the CpxR protein and both
the csgD and csgB promoters. The CpxRA pathway is
induced in response to damage of envelope proteins, such as during
exposure to elevated pH (10, 33), and to alteration of the
inner membrane lipid composition (11, 31). Activation of
the Cpx pathway results in the production of factors involved in
protein folding and degradation, such as the two
peptidyl-prolyl-isomerases PpiA (9) and PpiD
(12), DsbA, and the protease DegP (9). However, CpxR also represses motility and chemotaxis genes
(14) and is involved in regulation of P pili
(26), indicating that the Cpx pathway could play a role in
other cellular processes as well. We propose a new model that
integrates the complex regulatory networks controlling curli biogenesis.
MATERIALS AND METHODS
Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
TABLE 1.
E. coli K-12 strains and plasmids used
Genetic methods. Phage P1 transductions were carried out as described by Miller (32). The ompR234 mutation was transferred by using its genetic linkage (50% cotransduction) with malA, followed by screening of adherent transductants in 24-well microtitration plates. Transduction of the rpoS::Tn10 allele was obtained by selection on tetracycline plates and verification of the loss of catalase activity (29).
Enzyme assays.
-Glucuronidase specific activity in
toluene-treated samples was measured by spectrophotometrically
monitoring the hydrolysis of
p-nitrophenyl--D-glucuronide into
p-nitrophenol at 405 nm (4). Specific activity
was expressed as units per milligram of protein, where 1 U corresponds
to 1 nmol of product liberated per min (40).
-Galactosidase activity was measured by following the degradation of
o-nitrophenyl--D-galactoside into
o-nitrophenol, which absorbs at 420 nm (32).
Specific activity was expressed as nanomoles of product liberated per
minute per milligram (dry weight) of bacteria. A minimum of four
independent assays were performed, and the results were averaged to
obtain the indicated activities. Error bars indicate the standard deviation.
Adhesion and biofilm formation assays. Determination of biofilm thickness in microtiter plates was carried out as described by Dorel et al. (15). The ratio between surface-attached and unattached bacteria was estimated by measuring the optical density at 600 nm (OD600). At least three independent assays were performed and averages were calculated. To determine initial attachment to a solid surface, we used the sand column method described by Simoni et al. (48). Bacteria were grown in the appropriate medium, harvested, washed, and resuspended in phosphate-buffered saline (PBS) to an A280 of 1.0 (corresponding to ca. 5 × 108 bacteria); however, the adhesion properties of bacteria in the sand column assay depend strictly on the growth medium used (48) (data not shown). The suspension was loaded onto a fine sea sand grain column (9 g of sand). The bacterial concentration in the fractions collected at the column outlet was determined spectrophotometrically and used to calculate the percentage of attaching bacteria. The accuracy of spectrophotometric measurements was confirmed by direct plate counts (data not shown). Microscopic analysis of the column sand grains shows that bacteria attach as single cells in the conditions used in our experiments and that the cell sizes of the different strains are comparable (data not shown).
Construction of a csgD::uidA fusion. To obtain a csgD::uidA chromosomal fusion, a 3,642-bp DNA fragment corresponding to the whole csg region was amplified by PCR from MG1655 chromosomal DNA as the template and by using primers C1 (5'-CGA ATA ATC TTG CGG TCG ACA AGC AGG-3') and C2 (5'-GAA AGT GCC GCA AGG AGC TCT AAC G-3'), which contain, respectively, SalI and SacI restriction sites (italic sequences). The PCR fragment digested with SalI and SacI was cloned into the corresponding sites of the vector pBC (Stratagene) to give pBCcsg. The 3.8-kb SmaI fragment containing a uidA-kan cassette (pN496) (25) was cloned into the unique EcoRV site of the csgD gene carried by pBCcsg, producing pBCcsgD::uidA. The correct csgD::uidA orientation was confirmed by restriction digestion.
Integration of the plasmid into the chromosome was obtained by marker exchange mutagenesis, as described by Roeder and Collmer (43), followed by P1 transduction into the curli-producing strain PHL744. Transductants were selected for their inability to produce curli (white colonies on Congo red indicator [CFA] plates) and kanamycin resistance.-Glucuronidase (the product of the
uidA gene) specific activity was measured as described above.
Primer extension analysis of transcript. Total RNA was isolated from E. coli cells grown to an OD600 of 0.2 (2.5 × 109 CFU/ml) or 1 (1010 CFU/ml) in M9/glucose medium at 28°C, as described by Sambrook et al. (47). For csgB transcript analysis, we used the 5'-CCCAGGCGCACCCAGTATTGTT-3' primer, which anneals to the coding strand between 117 and 139 nucleotides downstream of the csgB gene transcription start. The sequence of the primer used for csgD transcript analysis was 5'-AAGATTTAGTGATCAACAATAATG-3', annealing to nucleotides +181 to +203 of csgD. The primers were labeled with the fluorescent dye IRD-800 at the 5' end. Extension products were run on a sequencing gel and densitometrically analyzed in a 4000L automated sequencer (Li-Cor Inc., Lincoln, Neb.).
Overproduction and purification of the OmpR and OmpR234 proteins. The coding regions of OmpR and OmpR234 were amplified by PCR from chromosomal DNA of, respectively, MG1655 and MG1655 ompR234 strains and by using the primers R1 (5'-AGTACAAACCATGGAAGAGAACTAC-3') and R2 (5'-CTTCGTACGCGAAAGCTTTATTAAACTG-3') carrying, respectively, an NcoI and an HindIII site (italic sequences). The presence of an NaeI cutting site in the ompR234 but not in the ompR amplified fragment (50) was checked. The 864-bp NcoI-HindIII fragments were then subcloned in the NcoI and HindIII unique sites of the plasmid cloning vector pKK233-2. The resulting plasmids, pKKompR and pKKompR234, contain fusions of the ompR and ompR234 start codons with the ATG start codon of the strong regulated trc promoter, placing ompR and ompR234 under the transcriptional and translational regulatory signals of the trc promoter.
Plasmids pKKompR234 and pKKompR were introduced by transformation into strain PHL694, which carries a chromosomal ompR null mutation (see Table 1). Three hours after isopropylthiogalactopyranoside (IPTG) induction, cells were harvested and suspended in 20 mM Tris-HCl (pH 7.4), 0.5 mM phenylmethylsulfonyl fluoride, 1 mM EDTA, and 1 mM dithiothreitol (DTT). Crude protein extracts were obtained by disrupting bacteria at 138,000 kPa in a French pressure cell (Aminco). OmpR and OmpR234 proteins were purified by fast protein liquid chromatography (FPLC) on DEAE-cellulose column chromatography as described by Jo et al. (27). The purification of the OmpR protein yielded protein of 85% purity, as judged by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE). Protein concentrations were determined with the Bio-Rad protein assay kit.Overproduction and purification of the CBP-CpxR protein. To construct the plasmid producing the calmodulin binding peptide (CBP)-CpxR fusion protein, a 700-bp region corresponding to the CpxR open reading frame was amplified by PCR using E. coli MC4100 genomic DNA as the template with primers cpx+ (5'-TATTTAAACCATGGATAAAATC-3') and cpxrev (5'-CTATCATGAAGCTTAAACCATC-3') carrying, respectively, an NcoI and an HindIII site. The amplified DNA was digested with NcoI and HindIII and subcloned into the corresponding restriction sites of pCAL-n (Stratagene), generating pCAL-n-cpxR, which was subsequently introduced into the BL21 strain. The pCAL vectors use the T7 lac promoter configuration and contain a copy of the lacI gene. On induction with 1 mM IPTG, the lacUV5 promoter was derepressed, allowing overexpression of T7 RNA polymerase and expression of the T7-promoted cpxR gene. Crude protein extracts were obtained as described above and incubated overnight with calmodulin affinity resin in CaCl2 binding buffer, according to the instruction manual (Stratagene). Washes and elution were performed as recommended by the manufacturer.
Gel retardation assay.
DNA probes containing either the
csgDEFG promoter region (128 to +12) or the same region
with deletions of the putative OmpR- and CpxR-binding sites, were
obtained by PCR amplification from pCSG4. For the wild-type promoter,
PCR amplification was performed by using primers D1
(5'-CCAAATGTACAAGCTTTCTATCATTTC-3') and D2 (5'-GGATTACATCTGATTTCAATCTAGCC-3'). For the promoter with a
deletion of the putative OmpR binding site, a 119-bp fragment was
amplified using two primers, D1 and D3
(5'-GGATTACATCTGATTTCAATCTAGCCATTACAAATCTTAAATCAAGTGTTCTCGTTATATTAAAATG-3'). The D3 primer sequence is identical to the sequence of D2 for its first
25 bases and then contains a 20-bp deletion corresponding to the
putative OmpR binding site (Fig. 1). The sequence of the deleted
operator was checked by automated sequencing using D1 primer (Genome
Express France). A DNA probe containing the csgB promoter
region was obtained by PCR amplification from pCSG4 using D4
(5'-CTGTCTGAAGCTTTTTGATAGCGGAAAACGG-3') and D5
(5'-CACCCTGGACCTGGTCGTACATTTAA-3') primers. All three
operator fragments were digested with HindIII and then
32P-labeled by the Klenow fragment of DNA polymerase.
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RESULTS |
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Top
Abstract
Introduction
Materials and Methods
Results
Discussion
References
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OmpR protein promotes curli production by activation of
csgD promoter.
As shown in a previous report, the
presence of the ompR234 allele favors biofilm formation by
stimulating the expression of the csgBA operon and the
production of curli (50). However, it is not clear if
OmpR234 activates csgBA by binding to the csgB promoter or in an indirect fashion. Regulation of the csgBA
operon is fairly complex and involves the transcription activator CsgD, the product of the first gene in the csgDEFG operon
(3, 44). In order to establish whether CsgD alone is
sufficient for csgB activation, we overexpressed the CsgD
protein in an OmpR
strain.
49.5 relative to the transcriptional start site of csgD
(Fig. 1). This sequence,
5'-GTTACATTTA/GTTACATGTT-3', closely resembles the consensus for OmpR binding sites proposed by
Harlocker et al. (21), and its location would be
consistent with possible OmpR-RNA polymerase interactions at the
csgD promoter.
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118 to
+12) and the promoter region with a deletion of the putative OmpR
binding site. Gel retardation assays shown in Fig.
2A and 2B clearly demonstrate that both
OmpR and OmpR234 can bind wild-type csgD but fail to
interact with the promoter region with a deletion of the putative OmpR
box (Fig. 2C). From these experiments we conclude that the OmpR protein
interacts directly with and activates transcription from the
csgD promoter by binding to a 20-bp region centered at
49.5. Identical concentrations of OmpR failed to retard a DNA
fragment encompassing the csgB promoter (173 to +88; data
not shown), strongly suggesting that the csgD promoter is
the main target for OmpR regulation. Similar results were obtained in
bandshift assays with purified His6-OmpR and
His6-OmpR234 proteins (data not shown).
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ompR234 mutation allows biofilm formation at high
osmolarity by transcriptional enhancement of both csg
operons.
Expression of the csgD promoter in E. coli and of agfD in S. enterica serovar
Typhimurium is negatively affected by high osmolarity, consistent with
a role of the envZ/ompR system in their
regulation (35, 44). We tested the effects of the osmotic
conditions in the growth medium on biofilm formation and on
csgDEFG and csgBA expression levels. Osmolarity
variations were achieved by adding increasing amounts of osmolytes such
as NaCl (Fig. 3) and sucrose (data not
shown) to low-osmolarity minimal M63/2 medium. The results of increased
medium osmolarity on biofilm formation, measured as crystal violet
staining and from OD600 measurement of cells attached to microtiter plates, are shown in Figure 3.
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ompR234 mutation increases initial adhesion to solid
substrates in a stationary-phase-dependent but
rpoS-independent manner.
The formation of biofilm
takes place in several steps, which include initial attachment to a
solid surface, formation of a microcolony, and differentiation into a
complex structure (7). To test if the ompR234
mutation affects initial adhesion, we used the sand column system
described by Simoni et al. (48), which provides a simple
and direct method to measure attachment. Cells grown overnight in
minimal medium were loaded onto sand columns after being washed and
resuspended in PBS. As shown in Fig. 5, for strain MG1655 (wild-type ompR) grown to the stationary
phase, only around 30% of the cells attached to the column. This shows that strain MG1655 has a weak ability to adhere to sand grains. In
sharp contrast, for the otherwise isogenic strain MG1655
ompR234, more than 70% of the cells adhered to the sand
grains.
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S, the
stationary-phase-induced alternative factor of RNA polymerase, in
both MG1655 and MG1655 ompR234. The newly produced EB1.3
(rpoS) and EB2.16 (rpoS ompR234)
strains were tested for adhesion to sand columns. Inactivation of the
rpoS gene did not significantly affect the ability to adhere
for either ompR234 or wild-type ompR strains
(Fig. 5). The lack of effects following rpoS inactivation was surprising, since rpoS is a positive regulator of the
curli operons in some E. coli strains and in
Salmonella (3). Thus, we tested the effects of
rpoS inactivation on csgDEFG and csgBA transcription by primer extension experiments.
As shown in Fig. 6, rpoS
inactivation stimulated both csgD and csgBA
transcription in the stationary phase (Fig. 6, lane 4). Expression of
the csgBA operon was detected in the stationary but not in
the exponential phase, consistent with the results obtained in the
adhesion experiments (Fig. 5). In contrast, higher levels of
csgD transcription were detected in the exponential phase,
earlier than the transcription from its target promoter csgBA, suggesting that CsgD might be subjected to
posttranscriptional regulation and might possibly regulate its own
expression. Experiments with luciferase reporter genes under the
control of either the csgB or the csgD promoter
confirmed the results of primer extension experiments (E. Brombacher
and P. Landini, unpublished data). Our results show that
S is not required for transcriptional
induction of csgB and csgD promoters and suggest
that E70 is preferably used upon entry into
the stationary phase in the ompR234 strain.
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Negative regulation of curli operon by transcription regulator
CpxR.
In a previous paper, we have shown that the two-component
regulatory system CpxA/CpxR negatively affects csgBA
expression (15). Both the csgD and
csgBA promoters display sequences with high similarity to
the proposed binding site for CpxR (36). To investigate
the possibility of a direct role of CpxR in curli regulation, we
purified CpxR and assayed its specific DNA binding to the
csgD and the csgB promoter regions. Indeed, CpxR
binds specifically to both the csgD and the csgB
promoters (Fig. 7), showing that CpxR is
directly involved in the downregulation of curli expression.
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pcsgD. D1D3 probe without protein (lane
5) and with 300 nM CBP-CpxR (lane 6). (B) pcsgB. D4D5
probe without protein (lane 7); D4D5 probe with 300 nM CBP-CpxR (lane
8); same as lane 8 but challenged with 100-fold excess of calf thymus
DNA unrelated binding site (lane 9); same as lane 8 but with a 40-fold
excess of competing unlabeled D4D5 DNA (lane 10). Arrows C,
CBP-CpxR-DNA complex.
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RS88[cpxP-lacZ]) (lane 1) and PHL1152 (MC4100
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DISCUSSION |
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
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The genetic organization and transcriptional regulation of curli-related operons csgBA and csgDEFG are highly conserved in E. coli and S. enterica serovar Typhimurium strains (44). Expression of these genes responds to different environmental signals and is positively regulated by ompR and rpoS (44). However, in strains MC4100 and MG1655 of E. coli, the expression of the csgBA genes is negligible despite the presence of functional ompR and rpoS genes. In a previous report, we showed that a G-to-T mutation in the ompR gene (ompR234 mutation), corresponding to a leucine-to-arginine substitution at position 43 of the OmpR protein, resulted in an increase in the expression of curli (50). In this paper, we have shown that increased curli production is mediated by OmpR234-dependent stimulation of transcription of the csgDEFG operon (Table 2). Increased production of the CsgD transcription activator results in activation of the csgBA operon; when CsgD is expressed independently of OmpR, the latter becomes dispensable for activation of the csgB promoter (Table 2).
Both genetic and biochemical evidence shows that OmpR binds to an
imperfect direct repeat (5'-GTTACATTTA/GTTACATGTT-3')
centered at position 49.5 relative to the transcriptional start site
of the csgD promoter (Table 2 and Fig. 1 and 2). This
sequence is very similar to the consensus for OmpR binding sites
proposed by several groups (21, 22, 37). In vitro binding
experiments show that both the OmpR wild-type and OmpR234 mutant
proteins can recognize this sequence, although OmpR234 displays a
slightly higher binding affinity than the wild-type protein, which
might be enough to improve activation of csgD transcription.
The OmpR234 protein might be better able to counteract the negative
regulation of csgD by the CpxR protein (15)
(Fig. 7A), either by competing for the same binding site or by inducing
structural changes in the csgD promoter region.
Alternatively, the leucine-to-arginine substitution in the OmpR234
protein (L43R) might improve its interaction with RNA polymerase.
The observation that, unlike the situation in other E. coli
strains and in S. enterica serovar Typhimurium (2,
44), the rpoS gene is no longer necessary for
csgD transcription in the ompR234 strain (Fig. 6)
might suggest that the L43R substitution results in a better
interaction of OmpR with 70 RNA polymerase.
Interestingly, a single-nucleotide substitution in the putative binding
site for OmpR in the S. enterica serovar Typhimurium
agfD promoter region also makes agfD
transcription rpoS independent (45).
The ompR gene is part of the ompR/envZ two-component regulatory system that senses osmolarity; thus, we investigated the possibility that biofilm formation might be regulated by osmolarity via the ompR/envZ pathway. Increasing NaCl or sucrose concentrations in the growth medium resulted in the inhibition of biofilm formation and decreased in vivo expression of the csgDEFG and csgBA operons, as measured with csgD::uidA and csgA::uidA chromosomal fusions (Fig. 3 and 4). Inhibition by NaCl or sucrose was observed in laboratory strains as well as in clinical isolates (Fig. 3), suggesting that osmolarity regulation of biofilm formation is broadly conserved in E. coli strains. Inhibition of curli production in the MG1655 strain PHL565 was observed at NaCl concentrations lower than those present in commonly used growth media (Fig. 3 and 4), providing an explanation for the previously reported lack of curli expression in laboratory strains (6, 19, 34).
Increasing osmolarity of the growth environment activates the sensor protein EnvZ and leads to increased phosphorylation of OmpR, resulting in enhancement of its DNA-binding ability (1, 13, 17, 24, 46). However, increased phosphorylation of OmpR results in activation of transcription only at certain promoters, such as ompC, while other genes, such as ompF, are downregulated. Repression of ompF transcription is due to the binding of phosphorylated OmpR (OmpR-P) to multiple binding sites, with the consequent inhibition of RNA polymerase binding to the ompF promoter (49). This mechanism might also apply to the regulation of csgD, although we were not able to identify multiple binding sites for OmpR (Fig. 1). It is possible that the negative regulation of csgD expression at high osmolarity is mediated by CpxR, since high osmolarity does indeed activate the Cpx pathway (Fig. 9). Control of cpxR expression by osmolarity would be consistent with the observation that increased osmolarity also results in a further decrease in the basal levels of csgBA transcription in a csgD mutant strain (Table 2).
Adhesion experiments using sand columns strongly suggest that the stage of biofilm formation positively affected by the ompR234 mutation is the initial adhesion to a solid surface (Fig. 5). Initial adhesion experiments also showed that the ompR234 mutation stimulates adhesion only in stationary-phase cells (Fig. 5), consistent with the simultaneous increase in csgB expression (Fig. 6). However, this stationary-phase-specific effect is not mediated by the master regulator RpoS, which, on the contrary, appears to negatively regulate csgBA expression in an ompR234 strain (Fig. 6).
We propose that negative regulation of csgB transcription by rpoS is due to rpoS-dependent transcription of cpxR (14). Interestingly, the CpxR protein binds both csgD and csgBA promoter regions, where it acts as a repressor (Fig. 7) (15). Lack of cpxR transcription in the rpoS-deficient strain would allow increased csgB transcription in the stationary phase. Our observations reiterate the importance of different regulatory networks in the regulation of the curli operon in E. coli and suggest that the CpxRA pathway plays a major role in the expression of virulence factors such as curli (this report) and P pili (26). The extreme complexity of the regulation mechanisms is likely to reflect the importance of finely tuning the expression of adhesion genes for survival of the bacterium in different environments.
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ACKNOWLEDGMENTS |
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We thank Sylvie Reverchon for critical reading of the manuscript, Valérie Gaubiac and Véronique Ramos for technical help, and also Valérie James for English corrections. We thank T. Silhavy for the gift of strains.
This work was partly supported by research grant 3100-058871 from the Swiss National Science Foundation and by a grant from the Centre National de la Recherche Scientifique (Réseau "Infections Nosocomiales").
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FOOTNOTES |
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* Corresponding author. Mailing address: Unité de Microbiologie et Génétique (CNRS ERS 2009), Institut National des Sciences Appliquées de Lyon, 11 avenue Jean Capelle, 69621 Villeurbanne Cedex, France. Phone: (33) 4 72 43 80 88. Fax: (33) 4 72 43 87 14. E-mail: dorel{at}insa-lyon.fr.
Present address: UMR CNRS 5557 Ecologie Microbienne,
Université Claude Bernard Lyon I, 69622 Villeurbanne Cedex, France.
Present address: Laboratoire d'Ingiénerie des
Systèmes Macromoléculaires, CNRS, Marseille, France.
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REFERENCES |
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Abstract
Introduction
Materials and Methods
Results
Discussion
References
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Journal of Bacteriology, December 2001, p. 7213-7223, Vol. 183, No. 24
0021-9193/01/$04.00+0 DOI: 10.1128/JB.183.24.7213-7223.2001
Copyright © 2001, American Society for Microbiology. All rights reserved.
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