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Infection and Immunity, February 2007, p. 745-752, Vol. 75, No. 2
0019-9567/07/$08.00+0     doi:10.1128/IAI.01163-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

Iron-Responsive Repression of Urease Expression in Helicobacter hepaticus Is Mediated by the Transcriptional Regulator Fur{triangledown}

Clara Belzer, Bart A. M. van Schendel, Ernst J. Kuipers, Johannes G. Kusters, and Arnoud H. M. van Vliet*

Department of Gastroenterology and Hepatology, Erasmus MC—University Medical Center, Rotterdam, The Netherlands

Received 25 July 2006/ Returned for modification 8 September 2006/ Accepted 2 November 2006


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Persistent colonization of mucosal surfaces by bacteria in the mammalian host requires concerted expression of colonization factors, depending on the environmental conditions. Helicobacter hepaticus is a urease-positive pathogen that colonizes the intestinal and hepatobiliary tracts of rodents. Here it is reported that urease expression of H. hepaticus is iron repressed by the transcriptional regulator Fur. Iron restriction of growth medium resulted in a doubling of urease activity in wild-type H. hepaticus strain ATCC 51449 and was accompanied by increased levels of urease subunit proteins and ureA mRNA. Insertional inactivation of the fur gene abolished iron-responsive repression of urease activity, whereas inactivation of the perR gene did not affect iron-responsive regulation of urease activity. The iron-responsive promoter element was identified directly upstream of the H. hepaticus ureA gene. Recombinant H. hepaticus Fur protein bound to this ureA promoter region in a metal-dependent matter, and binding resulted in the protection of a 41-bp, Fur box-containing operator sequence located at positions –35 to –75 upstream of the transcription start site. In conclusion, H. hepaticus Fur controls urease expression at the transcriptional level in response to iron availability. This represents a novel type of urease regulation in ureolytic bacteria and extends the already diverse regulatory repertoire of the Fur protein.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Urease is considered to be an important colonization factor for many pathogenic bacteria (8). The urease enzyme catalyzes the hydrolysis of urea into bicarbonate and ammonia. Both ammonia and bicarbonate are thought to contribute to acid resistance (27), whereas ammonia may also aid in injury of the surrounding epithelial cells (40) and serves as nitrogen source (52). In most bacterial species, expression of urease is strictly regulated in response to environmental or intracellular stimuli, such as substrate availability, growth phase, nitrogen status, or pH (1, 8, 11, 33).

Helicobacter species colonize the mammalian gastrointestinal and hepatobiliary tracts, resulting in chronic inflammation, which may progress to ulceration and carcinogenesis (21, 25). The murine pathogen Helicobacter hepaticus colonizes the intestine, bile ducts, and liver, and this can result in hepatitis, hepatic malignancies, and possibly cholesterol gallstones (22, 29, 50, 51). The urease enzyme is an important virulence factor in gastric Helicobacter species such as Helicobacter pylori and Helicobacter mustelae (2, 15, 34), but a contribution of urease in the development of H. hepaticus-associated disorders is yet to be demonstrated. We have previously demonstrated that urease expression in H. hepaticus is neither nickel or pH responsive nor growth phase regulated (5). This contrasts with the situation in H. pylori, where urease expression is controlled by nickel and pH at the transcriptional, posttranscriptional, and translational levels (1, 39, 43, 44). Although urease expression in H. hepaticus is lower than in H. pylori (5, 30), it is still relatively high compared to many other urease-positive pathogens (10). In view of the metabolic cost of urease expression and the importance of the enzyme in other bacteria, we hypothesized that urease expression in H. hepaticus is also regulated but may be responsive to a different environmental signal.

Iron is an important micronutrient and is required by almost all organisms (37). Iron is used as a cofactor of enzymes and participates in redox reactions. In the mammalian host, iron is mostly complexed into host proteins such as hemoglobin and ferritin, whereas at mucosal surfaces, iron availability is restricted due to chelation of iron by lactoferrin (37). Conversely, pathogenic bacteria often use iron restriction as an environmental cue for the expression of virulence factors such as hemolysins, toxins, and iron acquisition systems (38). Many of these iron-responsive virulence factors are under the control of the ferric uptake regulator (Fur) protein. Fur acts as a transcriptional repressor of iron-regulated promoters via iron-dependent DNA-binding activity (20). Under iron-replete conditions, Fur complexes with Fe2+ can recognize and bind to operator sequences (Fur boxes) in promoters. When iron is scarce, Fur dissociates from the DNA, allowing access of RNA polymerase to the promoter and transcription of the iron-regulated genes. The functions of Fur even extend outside iron metabolism, as the protein has also been reported to regulate oxidative stress resistance, acid resistance, and virulence factors, suggesting a key role for this protein in chronic colonization by pathogenic bacteria (7, 14, 16, 17, 23, 28).

In this study it is demonstrated that H. hepaticus uses iron as a signal for the regulation of urease expression and that this iron-responsive regulation of urease is mediated by the binding of the Fur protein to an operator sequence in the H. hepaticus ureA promoter. To our knowledge, this is the first demonstration of a direct role of Fur and iron in the regulation of urease expression; thus, this represents a novel function for Fur in the regulation of metabolic pathways and colonization factors.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Bacterial strains, plasmids, and growth conditions. H. hepaticus strain ATCC 51449 (22, 41) and its isogenic fur and perR mutants (this study) were routinely cultured at 37°C under microaerobic conditions (10% CO2, 5% O2, and 85% N2) on Dent agar (Oxoid) (44). Liquid growth was performed in brucella broth (Difco) supplemented with 0.2% (wt/vol) ß-cyclodextrins (Fluka) (BBC). Cultures were inoculated with a starting optical density at 600 nm (OD600) of 0.05 and were shaken at 40 rpm. For growth under iron-restricted conditions, 20 µM desferoxamine mesylate (Desferal; Sigma) was added to brucella broth before addition of the cyclodextrins. Iron-replete media were prepared by the addition of 100 µM FeCl3 (Sigma) to the iron-restricted medium. In case of transcriptional analyses (primer extension analysis), iron restriction was achieved by the addition of 2,2'-dipyridyl (Sigma) or 2,2'-dipyridyl and FeCl3 to final concentrations of 100 µM to an overnight culture of H. hepaticus, followed by incubation for 30 min at 37°C (14). The reason for the use of different chelators was that H. hepaticus RNA isolated after overnight growth in the presence of Desferal was of lower quality than RNA isolated from cells incubated with dipyridyl (data not shown) and could not be used for primer extension analysis, whereas incubation with 2,2'-dipyridyl was too short to result in phenotypic changes (protein profile and urease activity). Both chelators have been used previously for iron restriction in H. pylori and gave similar results for iron-responsive regulation of iron uptake and iron storage genes (6, 14, 16). Incubation of H. hepaticus with 2,2'-dipyridyl resulted in the absence of growth, probably due to the toxicity of the chelator (data not shown). Escherichia coli strains ER1793 and DH5{alpha} were grown aerobically in Luria-Bertani medium (36) at 37°C. When indicated, growth media were supplemented with chloramphenicol and ampicillin to final concentrations of 20 µg/ml and 100 µg/ml, respectively.

Urease enzyme assay. Urease enzyme activity was determined in freshly sonicated lysates by measuring ammonia production from the hydrolysis of urea, as described previously (44). The concentration of ammonia in the samples was inferred from a standard NH4Cl concentration curve. Enzyme activity was expressed as micromoles of urea substrate hydrolyzed per minute per milligram of protein.

Primer extension analysis. RNA was isolated from H. hepaticus with TRIzol reagent (Invitrogen), according to the manufacturer's instructions. Primer extension analyses were performed with the reverse primer HhureA-DIG (Table 1) essentially as described previously (17). The digoxigenin-labeled primer was annealed to 10 µg of total RNA from H. hepaticus strain ATCC 51449, and cDNA was synthesized after the addition of 5 units of avian myeloblastosis virus reverse transcriptase (Promega) and incubation for 1 h at 42°C. The cDNA product was separated on a 7% acrylamide-8 M urea sequencing gel and then blotted onto a nylon membrane (Roche), followed by chemiluminescent digoxigenin (DIG) detection (16). A sequencing reaction made with the same primer was used to determine the transcriptional start site.


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  		TABLE 1. Oligonucleotide primers used in this study

 
Construction of H. hepaticus fur and perR mutants. The fur gene of H. hepaticus strain ATCC 51449 was amplified with primers Hhfur-mutF1 and Hhfur-mutR1 (Table 1) and cloned in pGEM-T Easy (Promega), resulting in plasmid pCB1. The fur gene was subsequently interrupted by insertion of the chloramphenicol resistance gene from pAV35 (47) in the unique BsmI site, resulting in plasmid pCB2. The perR gene of H. hepaticus strain ATCC 51449 was amplified with primers HhperR-mutF1 and HhperR-mutR1 (Table 1) and cloned in pGEM-T Easy (Promega), resulting in plasmid pCB3. The perR gene was subsequently interrupted by insertion of the chloramphenicol resistance gene from pAV35 (47) in the unique HindIII site, resulting in plasmid pCB4. Both pCB2 and pCB4 were first introduced into E. coli ER1793, and plasmids isolated from E. coli ER1793 were subsequently used for natural transformation of H. hepaticus 51449 (5). Chloramphenicol-resistant colonies isolated were designated 51449fur and 51449perR, respectively. Two colonies derived from independent transformations were tested, and both colonies gave identical results in all experiments. Correct allelic replacement of the wild-type fur and perR genes with the interrupted version was confirmed by PCR using combinations of the primers Hhfur-outF1/Hhfur-outR1 and HhperR-outF1/HhperR-outR1, respectively (Table 1).

Protein analysis. H. hepaticus ATCC 51449 was grown for 24 h in iron-restricted and iron-replete BBC, centrifuged at 4,000 x g for 10 min at room temperature, and resuspended in phosphate-buffered saline, pH 7.4, to a final OD600 of 10. H. hepaticus cells were subsequently lysed by sonication for 15 s on ice with an MSE Soniprep 150 at amplitude 6. Proteins were separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) on a 10% (wt/vol) polyacrylamide gel and stained with Coomassie brilliant blue. Immunoblotting was performed after electrotransfer of proteins from a 10% SDS-polyacrylamide gel to a nitrocellulose membrane (Roche). The blot was subsequently probed with antibodies raised in rabbits to Helicobacter felis UreA or UreB (Intervet International BV, Boxmeer, The Netherlands) (5). Bound antibodies were visualized with swine anti-rabbit antibodies labeled with alkaline phosphatase (Promega), with BCIP (5-bromo-4-chloro-3-indolylphosphate) and Nitro Blue Tetrazoleum (Promega) used as a substrate (5).

Recombinant expression of H. hepaticus Fur protein. The fur gene was amplified from H. hepaticus ATCC 51449 with primers Hhfur-overF1 and Hhfur-overR1 (Table 1). The resulting fragment was digested with BamHI and PstI and ligated into pASK-IBA7 (IBA, Gottingen, Germany) to create pCB10. The wild-type sequence of the fur gene was confirmed by DNA sequencing. H. hepaticus Fur was expressed with an N-terminal Streptag, which has been used with the H. pylori Fur, NikR, and CrdR proteins and which does not influence DNA-binding activity (17, 18, 48, 49); therefore, the Streptag was not removed prior to use. The recombinant protein was purified as described in the manufacturer's instructions (48). The recombinant protein was over 90% pure, as determined by staining with Coomassie brilliant blue following electrophoresis on 12% SDS-polyacrylamide gels. Purified protein was dialyzed and stored in 50 mM Tris (pH 8.0), 100 mM NaCl, 1 mM MgCl2, 2 mM dithiothreitol (DTT), and 50% glycerol and was used without further purification for electrophoretic mobility shift and DNase I footprinting assays.

Electrophoretic mobility shift assays. DIG-labeled ureA promoter DNA was amplified with primers HhureAF1 and HhureA-DIG (Table 1). As a negative control, the promoter of the ksgA gene (HH1174) (41) was amplified with primers HhksgA-F1 and HhksgA-DIG (Table 1). The binding buffer used for the gel shift assays contained 24% glycerol, 40 mM Tris (pH 8.0), 150 mM KCl, 2 mM DTT, 600 µg/ml bovine serum albumin, and 1 µg denatured salmon sperm DNA. Gel shift assays were performed with 0.5 nM DIG-labeled ureA or ksgA promoter DNA, which was mixed with increasing concentrations of recombinant Fur protein and was subsequently incubated for 30 min in binding buffer at 37°C. Reactions were separated for 35 min at 200 V on an 8% polyacrylamide gel. The assay was performed in the presence of either 200 µM MnCl2 or 200 µM EDTA in binding buffer and PAGE buffer (2.5 M Tris and 0.19 M glycine). Gels were blotted onto a positively charged nylon membrane (Roche), followed by chemiluminescent DIG detection (44).

DNase I footprinting assays. DNase footprint assays were performed with 500 nM recombinant Fur protein and 50 nM DIG-labeled ureA promoter DNA or different concentrations of recombinant Fur in combination with 20 nM DIG-labeled ureA promoter DNA. Binding buffer consisted of 50 mM Tris (pH 8.0), 250 mM NaCl, 50 mM KCl, 5 mM DTT, 0.5% NP-40, 50% glycerol, and 100 µg/ml denatured salmon DNA in the presence or absence of 200 µM MnCl2. DNase I digestion was carried out by the addition of 0.25 units of DNase I (Promega) for 1 minute at room temperature, and the reaction was stopped by the addition of 2.5 M sodium acetate, 20 mM EDTA, and 10 µg denatured salmon DNA. Subsequently the DNA was ethanol precipitated and resuspended in 6 µl of loading buffer (0.05% bromophenol blue [Sigma] and 0.05% xylene cyanol [Merck] in deionized formamide [Sigma]), denatured, and separated on a 7% polyacrylamide gel containing 8 M urea. Gels were blotted onto a positively charged nylon membrane (Roche), followed by chemiluminescent DIG detection (44).


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Induction of H. hepaticus urease expression and activity by iron restriction. The effect of iron on H. hepaticus growth was determined by growing bacteria in BBC (standard conditions) or BBC supplemented with either desferoxamine mesylate (iron-restricted conditions) or desferoxamine mesylate and 100 µM FeCl3 (iron-replete conditions). Iron restriction resulted in reduced growth of H. hepaticus, and cells entered late log phase between 16 and 24 h of growth (Fig. 1A). In contrast, growth under standard conditions and iron-replete conditions saw cells in late log phase after approximately 24 h. At 24 h, cells were still viable, and examination of cells by Gram staining indicated that after 24 h in iron-restricted and iron-replete conditions the cultures still contained >95% bacillary forms of H. hepaticus. Changes in gene expression after 24 h are therefore likely to be a result of changes in iron availability rather than being growth phase dependent (16, 31); thus, we decided to use the 24-h time point for phenotypic analyses.


Figure 1
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  		FIG. 1. Induction of H. hepaticus urease transcription, expression, and activity under iron-restricted conditions. (A) H. hepaticus ATCC5 1449 was grown in standard BBC (gray squares), iron-restricted medium (white diamonds), and iron-replete medium (black triangles), and the OD600 was monitored over a time period of 36 h. The arrow indicates the 24-h time point, representing late log phase, which was chosen for comparison of iron-restricted and iron-replete conditions. (B) Urease activity in H. hepaticus is iron repressed, as measured after 24 h of growth in iron-restricted medium (white bar) or iron-replete medium (black bar). Each bar represents data from seven independent experiments; error bars denote standard deviations. An asterisk indicate a significant increase in urease activity after growth under iron-restricted conditions compared to growth under iron-replete conditions (P < 0.05 [Mann-Whitney U test]). (C) The increase in H. hepaticus urease activity under iron-restricted conditions is associated with increased expression of the UreA and UreB enzyme subunits, as shown by SDS-PAGE (left panel) and immunoblotting (right panel) with antibodies to H. felis urease. The left panel displays the relevant section of the protein profile of the lysates used for immunoblots, stained with Coomassie brilliant blue. Relevant marker sizes are indicated on the left. (D) Identification of the H. hepaticus ureA transcription start site by primer extension analysis using RNA purified from H. hepaticus ATCC 51449 grown under iron-restricted (–Fe) and iron-replete (+Fe) conditions. The sequence of the corresponding promoter region is displayed on the right, with the +1 residue and the –10 promoter sequence indicated. Note that the primer extension product displays iron-responsive repression of transcription.

 
Growth of H. hepaticus under iron-restricted conditions resulted in a twofold increase in urease activity compared to cells grown under iron-replete conditions (Fig. 1B). This iron-responsive urease activity resulted from increased expression of the urease subunit proteins UreA and UreB, as demonstrated by SDS-PAGE and immunoblot assay using both UreA and UreB antibodies (Fig. 1C). These findings contrast with nickel induction of urease expression in H. hepaticus, as this is mediated solely by increased enzyme activity, without additional UreA or UreB protein expression (5). Induction of urease activity and expression by iron restriction was mediated at the transcriptional level and mediated from the ureA promoter, as shown by primer extension analysis (Fig. 1D). The transcription start site of the ureA gene was identified to be the G residue 47 bp upstream of the ureA ATG start codon (Fig. 1D). There was only a single transcription start site, which displayed iron-responsive repression (Fig. 1D) similar to the regulation of expression and activity of H. hepaticus urease (Fig. 1B and C). Upstream of the transcription start site there is a TACAAT hexamer sequence identical to the –10 sequence of the H. pylori ureA promoter (12), which suggests that the H. hepaticus urease cluster is transcribed via the {sigma}80 sigma factor. Further comparison of the H. hepaticus ureA promoter sequence with that of H. pylori indicated the absence of the recently identified NikR binding sequence (13, 18, 19), consistent with the absence of nickel-responsive transcription of the urease genes in H. hepaticus (5).

Fur, but not PerR, mediates iron-responsive regulation of H. hepaticus urease. Since iron-responsive regulation of urease expression is mediated at the transcriptional level, this suggested the involvement of an iron-responsive regulatory protein. The H. hepaticus genome encodes two orthologs of such regulators: the peroxide stress regulator PerR (HH0942) and the iron-responsive regulator Fur (HH0893) (41). Both genes are predicted to be transcribed as a monocistronic mRNA (41); thus, mutation of these genes is unlikely to result in polar effects on surrounding genes. To determine whether Fur or PerR was involved in the iron-responsive regulation of urease expression, isogenic H. hepaticus fur and perR mutants were created by insertion of a chloramphenicol resistance gene.

Mutation of either the perR or fur gene did not significantly affect the growth of H. hepaticus, since growth of the mutants was similar to that of the wild-type strain under both iron-restricted and iron-replete conditions (data not shown). Mutation of the perR gene did not affect iron-responsive regulation of urease activity, and the resulting urease activities were similar to that of the wild-type strain (Fig. 2), while mutation of fur abolished regulation of urease activity at the tested conditions (Fig. 2). It should be noted that urease activity in the fur mutant does not show complete derepression compared to the wild-type strain under iron-restricted conditions; thus, it is possible that other regulatory mechanisms play a role in urease regulation in H. hepaticus.


Figure 2
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  		FIG. 2. Fur, but not PerR, mediates iron-responsive regulation of urease expression in H. hepaticus. Urease activity of H. hepaticus strain ATCC 51449 and the fur and perR mutants was assessed after 24 h of growth in iron-restricted medium (white bars) or iron-replete medium (black bars). Each bar represents data from three independent experiments for each strain; error bars denote standard deviations. An asterisk indicate a significant increase in urease activity after growth under iron-resticted conditions compared to growth under iron-replete conditions (P ≤ 0.05 [Mann-Whitney U test]); NS, not significant.

 
Fur binds to the H. hepaticus ureA promoter in a metal-dependent matter. To determine whether the Fur-mediated regulation of urease transcription in H. hepaticus is direct or indirect, the H. hepaticus Fur protein was expressed as recombinant protein in E. coli. Recombinant H. hepaticus Fur protein was mixed with DIG-labeled ureA promoter and tested by electrophoretic mobility shift assay in the presence and absence of metal. Fur displayed concentration- and metal-dependent binding to the ureA promoter region of H. hepaticus, as the ureA promoter fragment shifted in the presence of Fur and MnCl2 (Fig. 3, top panel), whereas this shift was not detected in the presence of Fur and EDTA (Fig. 3, middle panel). As a negative control, Fur protein was also incubated with the promoter region of the ksgA gene of H. hepaticus in the presence of metal; this did not result in a shift of the DNA fragment (lower panel), indicating that binding of Fur to the ureA promoter is sequence specific.


Figure 3
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  		FIG. 3. H. hepaticus Fur displays metal-dependent binding to the ureA promoter. Shown is an electrophoretic mobility shift assay using recombinant H. hepaticus Fur protein and DIG-labeled ureA promoter DNA (PureA). In the absence of the iron substitution manganese (+EDTA, middle panel), Fur is unable to complex with the ureA promoter region and a shift is not observed. Only in the presence of manganese (+MnCl2, top panel) is Fur able to bind to the ureA promoter region and cause a mobility shift (indicated as PureA + Fur). No shift is observed when the Fur protein is incubated with the promoter region of the ksgA gene (PksgA) in the presence of manganese (+MnCl2, bottom panel). The concentration of Fur is indicated above the lanes; the concentration of DNA was 0.5 nM ureA or ksgA promoter DNA.

 
Fur binds to an operator sequence in the H. hepaticus ureA promoter. The Fur-binding site in the ureA promoter was subsequently identified by DNase I footprinting. Addition of recombinant Fur resulted in the protection of a single region in the ureA promoter only in the presence of metal. Absence of either Fur or metal did not result in protection of the ureA promoter DNA (Fig. 4A), a finding consistent with the data obtained by electrophoretic mobility shift assays (Fig. 3). Fur-mediated protection of the region in the ureA promoter was concentration dependent (Fig. 4B). The protected sequence was located between positions –35 and –75 upstream of the ureA transcription start site (Fig. 4 and 5A), and the operator sequence contained a single sequence resembling a Fur box, consisting of three NAT(A/T)AT hexamers in an F-F-x-R formation (26).


Figure 4
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  		FIG. 4. Identification of the Fur operator sequence in the H. hepaticus ATCC 51449 ureA promoter. (A) DNase I footprinting assay performed in the presence and absence of the iron substitute MnCl2, using 50 nM DIG-labeled H. hepaticus ATCC 51449 ureA promoter DNA and 500 nM Fur. On the left side are indicated the positions relative to the ureA transcription start site. On the right-hand side is indicated the position of the protected region (located from –35 to –75 relative to the transcription start site). Note that a protected region is observed only in the presence of both Fur and MnCl2. (B) Binding of ureA promoter DNA is dependent on the Fur concentration. Increasing concentrations of Fur were mixed with 20 nM DIG-labeled H. pylori ATCC 51449 ureA promoter in the presence of 200 µM MnCl2. The concentration of Fur protein used is indicated above the lanes. Positions relative to the ureA transcription start site are indicated on the left; the protected region is indicated on the right.

 

Figure 5
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  		FIG. 5. (A) Schematic representation of the H. hepaticus ureA promoter region with the location and sequence of the Fur-binding site indicated. The proposed ribosome-binding site (RBS), –10 promoter sequence, and transcription start site (+1) are indicated. (B) Annotation of the proposed Fur box present in the Fur-bound sequence in the H. hepaticus ureA promoter region, according to the F-F-x-R hexamer consensus sequence. Residues indicated in bold type match the 5'-NAT(A/T)AT consensus hexamer sequence.

 

   DISCUSSION
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Urease is an important colonization and virulence factor for many pathogenic bacteria (8). In view of both the possible deleterious effects of uncontrolled ammonia production and the metabolic cost of urease production, expression and activity of urease are often controlled by specific environmental signals. These include substrate availability, nitrogen status of the cell, growth phase, and pH (1, 8, 11, 33). Helicobacter species are known to have some of the highest urease activities among ureolytic bacteria studied. This can be accompanied by urease protein expression reaching up to 10% of the soluble protein fraction in H. pylori (3). With the exception of pH, the aforementioned regulatory mechanisms of urease expression are absent in the few tested Helicobacter species (5, 9, 39). To date, only the molecular mechanisms governing urease activity in H. pylori have been reported, where urease expression is regulated at the transcriptional and posttranscriptional levels by nickel and pH (1, 44), with transcriptional regulation being controlled via the NikR and HP0166 (ArsR) regulatory proteins (13, 18, 35, 43, 45).

Urease activity of H. hepaticus is lower than that of H. pylori but still reaches high levels compared to many ureolytic bacterial species (5, 8, 10, 30). We previously demonstrated that the urease system in H. hepaticus is nickel responsive at the enzyme activity level (5) but not at the transcriptional level. The H. hepaticus genome encodes three orthologs of metal-responsive regulatory proteins that are all at different locations in the genome: the nickel-responsive regulator NikR (HH0352), the peroxide stress regulator PerR (HH0942), and the iron-responsive regulator Fur (HH0893) (41). The absence of nickel-responsive transcriptional regulation of urease in H. hepaticus (5) already suggested that NikR is not involved in urease regulation. Since the regulatory repertoire of H. hepaticus is relatively limited (41), we hypothesized that one of the other metal-regulatory proteins could be responsible for urease regulation in H. hepaticus. Here it is reported that H. hepaticus uses iron restriction as a signal for the induction of urease transcription, expression, and activity (Fig. 1); that PerR is not involved in urease regulation (Fig. 2); and that the iron-responsive transcriptional regulator Fur directly mediates this process (Fig. 2 to 4). Both fur and perR are predicted to be transcribed as a monocistronic mRNA; thus, the observed phenotype of the fur mutant is unlikely to result from polar effects of the insertion of the chloramphenicol resistance cassette. In addition, both the fur and perR genes are surrounded by genes annotated as hypothetical proteins, without any predicted function in urease activity or metal metabolism (41).

Fur was first identified in E. coli as the repressor of iron acquisition systems and was subsequently shown to be a DNA-binding protein that requires iron as a cofactor (20). In recent years it has been shown that the regulatory role of Fur extends beyond iron uptake systems, since Fur is involved in the regulation of many cellular processes (20). In several bacterial species, Fur may regulate oxidative stress defense genes, acid resistance, and metabolic routes (often via small RNA molecules) (28, 42). To date, Fur has not been shown to directly mediate urease regulation. There is a single report of a Fur box in front of a (silent) urease gene cluster in enterohemorrhagic E. coli strain EDL933, and mutation of fur resulted in decreased urease activity in a different enterohemorrhagic E. coli isolate (24). However, direct involvement of Fur in urease regulation in E. coli has so far not been conclusively demonstrated; thus, it remains possible that the observed phenotypes in E. coli are not mediated by Fur but are indirectly regulated via a cascade or through polar effects. Therefore, our study remains to our knowledge the first report of a direct role of Fur in regulation of a urease gene cluster.

The Fur protein has also been described to be involved in acid resistance for several bacterial species, including Salmonella enterica serovar Typhimurium and H. pylori (7, 23). Urease expression of H. pylori is also acid responsive, and urease activity is required for colonization of the stomach (15, 34). Although it is tempting to suggest a role for Fur in the acid resistance of H. hepaticus, it was previously shown that the urease system of H. hepaticus is not acid responsive and that urease activity does not allow survival at low pH (5).

H. hepaticus urease activity is not completely repressed by iron, as even under iron-replete conditions there is urease activity (Fig. 1A), and the fur mutant does not show a complete derepression of urease activity (Fig. 2). This could be due to a role of other regulatory systems in urease expression, as in H. pylori (35). Alternatively, binding of Fur may result in only incomplete or intermediate repression. The region protected from DNase I digestion in the footprint assay is located from –35 to –75 and thus is located just on the edge of the canonical {sigma}80 promoter sequence of the ureA promoter. This may be an indication of the presence of a binding site for an additional regulatory system upstream of the ureA promoter, as has been described for NikR and ArsR in the H. pylori ureA promoter (13, 18, 35). Alternatively, binding of Fur may only reduce but not completely block access of RNA polymerase. However, it should be noted that urease activity is only an indirect readout and is subject to regulation at the posttranscriptional or posttranslational level (5). The demonstration of sequence-specific binding of Fur to the ureA promoter (Fig. 3 and 4) confirms the direct role of Fur in urease regulation in H. hepaticus. The ureA operator sequence bound by Fur contains a putative Fur box (Fig. 5B) which matches well with the proposed consensus sequence (20, 26, 42).

The known environmental signals used for regulation of urease expression or activity are all somehow involved in the urease reaction or downstream processes, be it enzyme cofactor, substrate availability, nitrogen status, or pH. The link between iron and urease is not directly apparent, as iron is not involved in any of these pathways, with the exception of iron-responsive regulation of ammonia-producing enzymes in H. pylori (46). However, iron restriction is often used by pathogenic bacteria as a signal for entering the host and is a well known signal for regulation of virulence gene expression (32). The sites of colonization of H. hepaticus are the murine intestine and liver, and both environments are thought to be iron limited (22). In the intestine, H. hepaticus has to compete for iron with many other bacteria, including the resident flora. In contrast, the liver is thought to be a rather sterile environment containing high concentrations of bile and urea (4). Bile components can chelate iron, and this may make the hepatobiliary tract also an iron-restricted site of colonization for H. hepaticus (4). Increased urease expression may help H. hepaticus to overcome such detrimental conditions through as-yet-unknown mechanisms. Similarly, iron restriction functions as a signal for increased expression of the ammonia-producing enzyme amidase of H. pylori (46), but the physiological role of this regulatory response is also still unknown. It is therefore conceivable that a low level of iron can serve as an indirect signal for encountering unfavorable conditions requiring additional urease activity. Such increases in urease activity may help the bacteria to colonize their niche, for instance, through competition for nitrogen sources with other bacteria.

In conclusion, H. hepaticus uses iron and Fur for regulation of urease expression, a type of regulation not previously described for a urease system in ureolytic bacteria. These findings are another example of the versatility of the members of the genus Helicobacter, extending the possibilities of their relatively limited regulatory repertoire by using well known regulatory proteins such as Fur to mediate the expression of novel target systems.


   FOOTNOTES
 
* Corresponding author. Mailing address: Department of Gastroenterology and Hepatology, Erasmus MC—University Medical Center, 's Gravendijkwal 230, 3015 CE Rotterdam, The Netherlands. Phone: 31-10-463-5944. Fax: 31-10-463-2793. E-mail: a.h.m.vanvliet{at}erasmusmc.nl. Back

{triangledown} Published ahead of print on 13 November 2006. Back

Editor: V. J. DiRita


   REFERENCES
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
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Infection and Immunity, February 2007, p. 745-752, Vol. 75, No. 2
0019-9567/07/$08.00+0     doi:10.1128/IAI.01163-06
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