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1 Department of Restorative Dentistry, University at Buffalo, School of Dental Medicine, Buffalo, NY 14214, USA
2 Department of Periodontics and Endodontics, University at Buffalo, School of Dental Medicine, Buffalo, NY 14214, USA
Correspondence
Violet I. Haraszthy
vh1{at}acsu.buffalo.edu
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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; Zambon, 1994) as well as sometimes life-threatening extra-oral infections such as endocarditis (Kaplan et al., 1989; Overholt, 1966; Townsend & Gillenwater, 1969). This micro-organism possesses a diverse and rather impressive array of virulence factors (Baehni et al., 1981; Christersson et al., 1987; Helgeland & Nordby, 1993; Meyer & Fives-Taylor, 1993; Mintz & Fives-Taylor, 1994, 1999). While it is not known which of these is involved in the pathogenesis of periodontitis, A. actinomycetemcomitans, like other successful pathogens, must be able to induce or repress the production of virulence factors in response to the environment. The mechanisms by which A. actinomycetemcomitans responds to environmental cues and the genes/proteins regulated are largely unknown.
Iron is essential for bacterial growth both as a nutrient and as a catalyst in the formation of hydroxyl radicals (Palyada et al., 2004). However, iron is tightly sequestered in human hosts in order to inhibit bacterial growth and to prevent oxidative damage. Consequently, micro-organisms have evolved complex systems to efficiently capture iron. Importantly, in order to avoid toxicity, micro-organisms achieve an effective iron homeostasis by tightly regulating the expression of genes encoding proteins involved in iron acquisition and metabolic response to iron availability (Ratledge & Dover, 2000).
In many bacteria, the Fur protein controls the transcription of these genes (Escolar et al., 1999; Hantke, 1981). The main function of the Fur protein is to repress iron acquisition genes under conditions of iron sufficiency and to induce iron transport systems under conditions of iron restriction. Under iron-rich conditions, Fur can form a complex with Fe2+ that binds to specific consensus sequences (‘Fur box’) in the promoter of iron-regulated genes, leading to transcriptional repression (Bagg & Neilands, 1987). Under low-iron conditions, Fur dissociates from the promoter, allowing gene transcription. Therefore, when most Fur-regulated genes are repressed, specific genes such as those involved in iron storage or stress responses may be activated. There is growing evidence that Fur and Fur-like repressor proteins also control the expression of virulence determinants in many bacterial species (Litwin & Calderwood 1993; Ochsner et al., 1995; Prince et al., 1993). In Escherichia coli, for example, the Fur protein transcriptionally regulates more than 90 genes (Escolar et al., 1999). Consequently, identification of genes whose expression is controlled by Fur may lead to the discovery of as yet unknown bacterial virulence factors.
We have cloned and characterized the fur gene in A. actinomycetemcomitans (Haraszthy et al., 2002) – one of the first global regulatory genes identified in an oral pathogen. Current research in our laboratory is focused on identifying Fur-regulated genes in A. actinomycetemcomitans. While some of these will undoubtedly be involved in routine iron metabolism, others may control the expression of virulence factors. In this study, Fur titration assay (Furta) was used to detect Fur-regulated genes. In addition, a fur mutant A. actinomycetemcomitans strain was created by selection for manganese resistance to define the role of Fur and to confirm genes controlled by this protein.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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. A. actinomycetemcomitans was routinely cultivated from frozen stock on tryptic soy blood agar supplemented with 0·5 % (w/v) yeast extract (Difco) and 5 µg haemin ml–1. A. actinomycetemcomitans strains were grown in trypticase soy broth supplemented with 0·6 % (w/v) yeast extract (TSBYE) in a humidified, 5 % CO2 atmosphere at 37 °C. Manganese-resistant A. actinomycetemcomitans strains were cultured with 10 mM MnCl2 added to the medium and allowed to incubate at 37 °C for 48–72 h in 5 % CO2. For some experiments, A. actinomycetemcomitans was grown under anaerobic conditions at 37 °C in an anaerobic chamber (Coy Laboratory Products). E. coli strains were cultured in Luria–Bertani (LB) broth (10 g Bacto-tryptone l–1, 5 g Bacto-yeast extract l–1, 10 g NaCl l–1, pH 7·5) at 37 °C in air. Iron-regulated lacZ operon fusions of A. actinomycetemcomitans were screened on MacConkey agar containing 30 µM FeSO4 (van Vliet et al., 2000). Antibiotics were added to LB medium as needed, at the concentrations indicated: 100 µg ampicillin ml–1, 45 µg kanamycin ml–1, 100 µg streptomycin ml–1 and 50 µg tetracycline ml–1. BHI medium with or without the iron chelator 2,2'-dipyridyl (Sigma), at a final concentration of 0·2 mM, was used to assess the effect of iron concentration on gene expression. Cells used to inoculate the iron-restricted medium were subcultured on the same medium to deplete internal iron reserves.
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DNA isolation.
Bacterial genomic DNA was isolated by an SDS-proteinase K method (Sambrook et al., 1989). Plasmid DNA isolation (Wizard Plus Minipreps, Promega), DNA restriction endonucleases, T4 DNA ligase and T4 polynucleotide kinase were used according to the manufacturers' specifications.
Preparation of bacterial cell lysates.
Whole-cell protein extracts were obtained from A. actinomycetemcomitans cells cultured aerobically in the presence of 5 % CO2 at 37 °C under iron-chelated or iron-rich conditions. Iron-chelated conditions were obtained by the addition of 2,2'-dipyridyl. Bacterial cells were harvested by centrifugation at 12 000 g for 10 min at 4 °C, and then washed three times in ice-cold PBS, pH 7·0. The cells were lysed (Standard Cell Lysis Buffer, Amersham) and stored in aliquots at –70 °C. The protein concentration was determined by hydrolysis/amino acid analysis or by the PlusOne 2-D Quant Kit (Amersham).
Furta.
Furta was performed as described by Stojiljkovic et al. (1994) and Tsolis et al. (1995). Chromosomal DNA from A. actinomycetemcomitans was partially digested with Sau3A1. Fragments between 3 and 6 kb were cloned into pUC18 previously digested with BamHI. E. coli strain H1780 was transformed with the library. The recombinants were distributed onto MacConkey agar containing 30 µM FeSO4, supplemented with 100 µg ampicillin ml–1. The cultures were incubated at 37 °C for 36 h and the Lac phenotype was recorded. E. coli containing plasmid pUC18 was used as a negative control and the cloned E. coli fur gene with its promoter region (a gift from Dr K. Hantke, Mikrobiologie II, Universität Tübingen, Germany) was used as a positive control. All experiments were repeated at least twice. The identity of the selected clones was determined by dsDNA sequencing using the dideoxy chain-termination procedure (Sanger et al., 1977). Plasmid DNA was prepared (Wizard Plus Purification System, Promega) and sequenced. A universal primer, a reverse universal primer and synthetic oligonucleotide primers were synthesized to obtain the entire gene sequence. The resulting nucleic acid sequences were compared to DNA databases (Altschul et al., 1990) and to annotated A. actinomycetemcomitans sequences (Roe et al., 1999).
Gel shift assay.
Gel shift assays were performed with minor modifications (DIG Gel Shift Kit, Roche) to determine if A. actinomycetemcomitans genes were able to bind A. actinomycetemcomitans Fur. Digoxigenin-labelled DNA fragments were generated by PCR using commercially obtained primers (Life Technologies Gibco-BRL) based on sequences obtained during this study and screened against the A. actinomycetemcomitans database. Primers for minC Forward 1 (TGAGGTTCGAGTTGCTGATT; –69 to –49) and Reverse 1 (CTTGAGCCATCACGAAATTCCAAG; +10 to –14), and Forward 2 (ATGACTGCGCTTTTCAATATAAAG; –153 to –1) and Reverse 2 (CACGAAATTCCAAGAAAAAGAG; –1 to –22), generated 79 bp and 153 bp fragments, respectively. Crude whole-cell extracts were prepared from cells grown in iron-rich medium as described by de Lorenzo et al. (1987). Whole-cell protein isolates were tested from the following: (1) Aawt expressing the native Fur protein; (2) E. coli H1780 with pUCAafur expressing the cloned Fur protein; (3) AaMnR strains expressing the mutated Fur protein; and (4) E. coli strain H1780 containing pUC18, which served as a negative control. Samples (approx. 20 ng) of bacterial cell extracts were mixed with the labelled DNA fragments in 5x binding buffer [100 mM HEPES, pH 7·6, 5 mM EDTA, 50 mM (NH4)SO4, 5 mM DDT, 10 % (w/v) Tween 20, 150 mM KCl, 100 µM MnCl2 and 1 µg poly d(I-C) nonspecific DNA]. The mixtures were incubated for 15 min at room temperature and then transferred to a 6 % nondenaturing polyacrylamide gel and electrophoresed. Following electrophoretic separation, the DNA–protein complexes were blotted by capillary transfer onto positively charged nylon membranes. The digoxigenin-labelled probes were detected by enzyme immunoassay using anti-digoxigenin-AP Fab fragments. The membranes were developed using the Genius Detection Kit (Roche).
Manganese-resistant A. actinomycetemcomitans.
Manganese selection was performed by a modification of the method of Hantke (1987). Approximately 108 cells of A. actinomycetemcomitans JP2 were distributed on TSBYE medium containing 10 mM MnCl2 and allowed to incubate at 37 °C for 48–72 h in 5 % CO2. The resulting colonies were restreaked onto the same medium to confirm the resistant phenotype. Genomic DNA was isolated from five MnR A. actinomycetemcomitans colonies. Each colony was obtained from separate plates, from experiments performed weeks apart. The entire fur gene was amplified (Expanded High Fidelity Plus PCR System, Roche) from each using fur-specific primers (5'-TATTTCTGTGATGCCATGGG-3'; 5'-TGCGCAATGATTAATGTCGCTA-3'), sequenced and compared to the wild-type fur gene.
Cloning of the A. actinomycetemcomitans fur mutant gene.
To compare the function of the mutant fur gene (AaMnR) with that of the wild-type fur gene (Aawt), E. coli strain H1780 was used. This strain has an undefined mutation of chromosomal fur and a fiu–lacZ gene that is regulated by the promoter-operator region of an iron-regulated outer-membrane protein gene, fiu (Schaffer et al., 1985). Under normal conditions, the mutated fur gene in E. coli H1780 permits unrestricted production of 
-galactosidase, indicated by a Lac+ (blue) phenotype on X-Gal media. However, when a plasmid containing a functional fur is introduced into strain H1780, there is derepression of fiu by Fur and reduced -galactosidase activity, indicated by a Lac– (white) phenotype on X-Gal media. Accordingly, strain H1780 was transformed with pUC18 containing the amplified fragments of either the mutant or the wild-type fur. -Galactosidase activity was assayed and calculated as described by Miller (1972). E. coli strains were cultured overnight in iron-rich LB medium and 108 cells were inoculated into M9 minimal medium supplemented with antibiotics and either 100 µM iron or 200 µM 2,2'-dipyridyl.
Western blot analysis.
Whole-cell protein extracts were prepared from bacteria grown to late exponential phase in TSBYE medium with and without 2,2'-dipyridyl. Thirty micrograms of protein extract from each strain was separated by electrophoresis on 12 % SDS-polyacrylamide gels containing prestained high-molecular-mass standards (Bio-Rad). Gels were stained with Coomassie blue or transferred onto nitrocellulose paper using a semi-dry blotting apparatus (Hoefer Scientific). Blots were blocked with 2 % (w/v) BSA in PBS with azide for a minimum of 1 h. The nitrocellulose transfers were reacted with antiserum to Pseudomonas aeruginosa Fur (Prince et al., 1993) diluted in blocking buffer. Positive reactions were detected using alkaline phosphatase. Protein concentrations were determined by the Bradford method with Coomassie protein assay reagent (Pierce).
Growth at acidic pH.
To determine whether mutation of the fur gene affected the growth of A. actinomycetemcomitans under acidic conditions, Aawt and AaMnR strains were cultured in broth under aerobic conditions ranging from pH 6·0 to 7·2 in increments of 0·1 unit (Clyne et al., 1995). The broth was adjusted to the desired pH with HCl. To ensure that all strains were in the same growth phase, the bacteria were grown to an OD600 of approximately 0·1 (SmartSpec 3000 spectrophotometer) in pH 7·0 broth and diluted in the test medium to an OD600 of 0·05. Bacterial growth was monitored by measuring the OD600 until stationary phase was reached, after which no increase in optical density was observed. The pH of the medium was monitored by pH electrode (accumet basic AB15, Fisher Scientific) and did not change by more than 0·5 unit during the course of the experiments.
To compare cell density at OD600 versus c.f.u. (per ml), aliquots of cultures were removed at various times during growth and plated in triplicate on TSBYE agar. All growth experiments were repeated at least three times with consistent results.
RNA isolation.
RNA was extracted from mid-exponential-phase cells of Aawt and AaMnR strains grown in iron-rich or iron-chelated medium (RNAqueous Extraction Kit, Ambion) according to the manufacturer's instructions. RNA yields were analysed spectroscopically.
Real-time quantitative PCR (RT-qPCR).
One-step RT-qPCR was performed to quantify gene expression. PrimerExpress software (version 2.0; Applied Biosystems) was used to design primers and probes. The primer sequences were examined (BLAST) to ensure that they were specific to the target gene. Primers had Tm values of between 58 and 60 °C. Probes did not overlap the primers and had Tm values 10 °C higher than the primers. The fluorescent reporter was attached to the 5' end of the probes. TaqMan probes had 5'-FAM and 3'-TAMRA modifications. The quencher was attached to the 3' end of the probes. The primers and probes used for real-time PCR (Applied Biosystems) are listed in Table 2. Reaction mixtures (20 µl) contained 20 ng template RNA, 1x TaqMan Master Mix, 0·25 units MultiScribe reverse transcriptase µl–1, 0·4 units RNase inhibitor µl–1, TaqMan probe (Applied Biosystems), at a concentration of 0·25 µM, and forward and reverse gene-specific primers at a concentration of 0·6 µM. All reactions were run in triplicate and repeated three times with freshly isolated RNA obtained from separately grown cells. Positive controls included (1) standard RT-PCR amplification with a known template producing a single amplicon of the expected size; and (2) amplification with a primer/probe set for afuA, a gene known to be Fur regulated. Negative controls included (1) amplification with a primer/probe set for 16S rRNA, which is not Fur regulated; (2) amplification with a reaction mixture without a template; and (3) amplification without the reverse transcriptase. ROX was used as an internal (passive) reference dye to normalize the signal of the reporter. Since the reference dye is added to each reaction in equal amounts, this controls for non-amplification-based variability in the results. Reaction plates were processed on an Applied Biosystems 7900HT Sequence Detection System. The reverse transcription reaction was performed at 48 °C for 30 min. The AmpliTaq Gold polymerase was activated at 95 °C for 10 min followed by 40 cycles of denaturation for 15 s at 95 °C and annealing and extension for 60 s at 60 °C. Amplification data were analysed with the ABI Prism SDS 2.1 software (Applied Biosystems). Relative quantification of gene expression was performed by the Ct method (Winer et al., 1999), with 16S rRNA expression serving as an endogenous control to normalize expression within each sample. By comparing the threshold cycle between the wild-type and mutant strain, the relative expression of a given gene could be determined. Statistical significance was determined by ANOVA and Fisher's multiple comparison analysis.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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), Furta utilizes E. coli strain H1780. This strain carries fhuF–lacZ, a Fur-regulated gene fusion sensitive to changes in Fur concentration. Fur-binding promoter regions carried on a high-copy-number plasmid can be identified by binding Fur in E. coli, preventing Fur binding to the lacZ construct located on the chromosomal DNA and allowing transcription of the -galactosidase gene. High levels of -galactosidase expression result in red E. coli colonies on MacConkey medium, indicating that the plasmid carries a Fur-binding fragment, whereas reduced -galactosidase expression results in a pale colony indicating that the plasmid does not carry a Fur-binding fragment (Stojiljkovic et al., 1994; Tsolis et al., 1995). Furta is a powerful tool for identifying Fur-regulated genes, but it is extremely sensitive to both iron and Fur concentrations. A. actinomycetemcomitans fur has 62 % homology to E. coli fur gene and the cloned A. actinomycetemcomitans fur gene was able to complement the fur mutation in E. coli (Haraszthy et al., 2002). However, the assay probably targets clones containing Fur boxes with a high degree of similarity to the E. coli Fur box since the E. coli Fur protein will bind those sites preferentially. This would suggest that mainly A. actinomycetemcomitans promoters recognizing E. coli Fur were detected in this study. In the future we intend to use more sensitive techniques such as proteomics to detect most of the Fur-regulated genes in A. actinomycetemcomitans.
Fifty potentially Fur-regulated clones were selected after screening 30 000 clones from an A. actinomycetemcomitans plasmid bank by Furta. When these 50 clones were sequenced, 14 were identified as containing potentially Fur-regulated genes and the remainder were identified as duplicates or false-positive clones. Clones were considered false positive if a ‘Fur box’-like sequence was not identified in their promoter region or the clone did not include the promoter region of the sequenced gene. Although it is possible that some of these genes might be Fur regulated by an unknown method they will be the subject of future studies. As shown in Table 3, several different types of genes were noted among the 14 potentially Fur-regulated genes. These include a previously described Fur-regulated gene, afuA, encoding a periplasmic ferric uptake protein belonging to the metal-transporting, binding-protein-dependent ABC systems (Willemsen et al., 1997) and another iron acquisition gene, hemA. A transferrin-binding protein gene, tbpA, also appeared to be Fur regulated. Three genes encoding outer-membrane proteins were identified, including a putative outer-membrane protein with no known function, (clone 2), a TonB-dependent outer-membrane siderophore receptor (clone 3) and a possible transcriptional regulator (clone 4). Putative genes encoding an adhesion molecule (clone 1), an ABC transporter (clone 6) and a possible permease (clone 7) were identified. A putative iron-transport gene similar to the yfe system of Yersinia pestis (Bearden & Perry, 1999) was also detected. Genes for various enzymes appear Fur regulated such as recO (DNA repair) and dgt (deoxyguanosine triphosphate triphosphohydrolase). A cell division inhibitor gene, minC, was also among the potential iron/Fur-regulated genes as was eno, encoding the glycolytic enzyme enolase, known to be important in the pathogenesis of streptococcal infections (Pancholi & Fischetti, 1998; Pancholi, 2001).
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; Berish et al., 1993; Funahashi et al., 2000); however, it is possible to produce fur mutants using manganese selection (Prince et al., 1993; Lam et al., 1994; Tolmasky et al., 1994, Benson et al., 2004). The MnR Fur mutants are generated using high concentrations of manganese. MnR A. actinomycetemcomitans were obtained by culturing strain JP2 on TSBYE medium containing 10 mM MnCl2. Comparison of the fur gene from five MnR colonies (AaMnR) with the wild-type (Aawt) revealed the same single point mutation in each. The colonies were picked from several different plates originating from separate cell cultures. There was a change in nucleotide 66 from C to T, changing leucine to isoleucine. Single point mutations and small insertions have been reported for other MnR fur mutants (Lam et al., 1994). The majority of the mutations are clustered at the N- and C-terminal regions of the protein. The AaMnR mutant contains a single point mutation in the amino-terminal domain where the helix–turn–helix domain is located, which may be important for DNA binding (Kolade et al., 2001), resulting in a nonfunctional Fur protein.
Manganese selection is a relatively specific method for isolating fur mutations in many species of bacteria (Brickman et al., 1995; Lam et al., 1994; Prince et al., 1993; Hantke, 1987). The molecular mechanism of manganese selection is not known, but it has been suggested that the ability of Mn2+ to activate Fur for DNA binding may be a source of manganese toxicity, perhaps by inappropriately repressing iron uptake functions (Bsat & Helmann, 1999). Another possibility is that the fur gene is highly sensitive to the mutagenic action of manganese (Prince et al., 1993). Manganese mimics iron by binding to the Fur protein to form a Fur–Mn2+ complex instead of the Fur–Fe2+ complex and repressing iron uptake genes. As a result, bacteria with the wild-type fur gene repress iron uptake systems and starve for iron in the presence of manganese, while fur mutants fail to repress iron-uptake systems and survive. The Fur protein in these mutants is thought to retain some function, explaining why this particular class of mutations is not lethal. It has been also suggested that some gene products that are required for viability may be produced only in the presence of Fur. It is possible that some of the iron acquisition genes are turned off in the AaMnR strain either directly by Fur or by another so far unidentified pathway to prevent the accumulation of iron inside the cells. It is also possible that the AaMnR strain survived iron starvation by acquiring a mutation which at least partially derepresses the iron uptake system (Thomas & Sparling, 1996). Alternatively, constitutive expression of all of the Fur-repressed genes may be detrimental to the growth of the bacteria because of the resulting intracellular iron overload. This type of explanation is supported by the work of Touati et al. (1995), who observed that fur mutants of E. coli produced iron overload, leading to oxidative stress and DNA damage. In our future studies we aim to create a fur null mutant A. actinomycetemcomitans strain to further define the Fur regulon. By comparing the two fur mutant strains we will be in a position to identify genes whose expression differs between these strains. Those genes could be of potential interest in defining the role of Fur in A. actinomycetemcomitans virulence.
The activity of the fur gene from AaMnR was tested in a fur-deficient E. coli strain H1780 by cloning it into pUC18. Since the fur gene of H1780 has been inactivated, the fiu–lacZ reporter gene is unregulated and constitutive. Introduction of a clone encoding an active Fur protein into H1780 re-establishes Fe-dependent regulation of the fiu–lacZ reporter gene, leading to a phenotype change. The fur gene cloned from the AaMnR strain was not able to regulate the fiu–lacZ reporter gene. The Lac+ (blue) phenotype of the clones indicated that the gene with the point mutation encoded a Fur protein that was unable to bind to the Fur-regulated promoter in E. coli. This indicates that the fur gene mutation in the AaMnR strain results in a non-functional Fur protein under aerobic conditions.
Effect of iron limitation on the growth of AaMnR
The effect of fur mutation on growth rate was examined by comparing the growth of AaMnR and Aawt in iron-rich and iron-chelated media. Similar to previously studied fur mutant strains (Litwin & Calderwood, 1994) the AaMnR strain grew more slowly than the Aawt strain under iron-rich conditions (Fig. 1). In the iron-chelated medium both strains had a very similar growth rate. This indicated that the Fur protein in the mutant strain is unable to respond to changes in environmental iron concentrations. The constitutive high-level expression of iron uptake systems in the AaMnR mutant may be an unneeded energy drain in the presence of sufficient iron and may, therefore, impede growth.
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). As expected, the wild-type A. actinomycetemcomitans strain responded to available iron by expressing significantly more Fur protein under iron-rich conditions than under iron-chelated conditions (Fig. 2). The AaMnR mutant strain was unable to respond to available iron and expressed equal amounts of Fur protein under both iron-rich and iron-chelated conditions (Fig. 2). These data are, therefore, consistent with the idea that the point mutation in the fur gene results in a non-functional or partially functional Fur protein, unable to respond to varying iron concentrations under the test conditions.
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) until stationary phase was reached. The Aawt strain and the AaMnR mutant had similar growth rates in pH 7·0 broth but the growth rate of the AaMnR strain at pH 6·8 was reduced compared to that of the parent strain. The wild-type A. actinomycetemcomitans adapted well to acidic conditions and was able to grow even at pH 6·0, indicating that it can withstand and adapt to acidic environments. The AaMnR strain grew poorly in acidic media, indicating that Fur is important in the ability of A. actinomycetemcomitans to adapt to an acidic environment. Relatively little is known about the mechanisms involved in the acid resistance of A. actinomycetemcomitans. The fact that this bacterium can persist and propagate in the oral cavity suggests that it is able to grow at mildly acidic pH. The ability to adapt to acidic environments could be important in the pathogenesis of A. actinomycetemcomitans infections since there are frequent episodes of high acidity in the human oral cavity (Bijlsma et al., 2002). Generally, bacteria growing at acidic pH respond by inducing changes in lipopolysaccharide composition (McGowan et al., 1998; Moran et al., 2002), increasing expression of chaperone-like proteins (Huesca et al., 1998), and expressing several genes at the transcriptional and/or translational level (Merrell et al., 2003; Ang et al., 2001). However, the exact role in acid resistance of many of these factors is largely unknown. Since iron is essential for growth, an environmental shift from high to low iron concentrations serves as an important regulatory signal to the cell. Because iron solubility is a function of pH, it is reasonable to speculate that Fur may sense acid indirectly in terms of iron availability. Therefore, Fur may play a role in adaptation to acidic environment by sensing acid stress and subsequently regulating the expression of some genes in an iron-independent fashion. The role of Fur in acid resistance should be the subject of future studies.
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, 1991). E. coli cells transformed with the pUCAaminC clone formed long chains, suggesting that the cells were unable to divide properly. This clone also grew slowly (data not shown).
A potential Fur box (AATAATTCTTTACTTATTA) was detected upstream of the minC start site. It contained 11 of 19 bases or 57·9 % identity to the E. coli Fur box. To test the potential regulatory effect of Fur, a 79 bp and a 153 bp fragment were amplified from the promoter region of the gene. Both of these fragments included the putative Fur box with flanking regions of various sizes. A. actinomycetemcomitans Fur retarded the mobility of the putative Fur consensus sequence in the gel mobility shift assays (Fig. 4). The Fur protein from the AaMnR strain or from E. coli containing pUC18 alone was not able to bind to the promoter region of the minC gene and retard its mobility. This experiment suggested that the gel shift seen in the binding reaction containing the promoter region of the minC gene resulted from the formation of a Fur protein–fur consensus probe complex, illustrating that minC does indeed contain a Fur box and that Fur binds to minC.
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found that A. actinomycetemcomitans afuA gene expression increased under iron-chelated conditions and that the fur gene promoter region contains a Fur box able to bind the Fur protein. We used this gene in our experiments as a positive control. Consistent with previous studies, our studies also showed increased afuA gene expression under iron-chelated conditions. There was an eightfold increase in the copy number of afuA transcripts in Aawt cultured under iron-chelated conditions compared to Aawt cultured under iron-rich conditions. In the AaMnR mutant, there was a five- to sevenfold increase in the copy number of the afuA transcripts compared to Aawt when both were cultured under iron-rich conditions (Fig. 5).
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), which would be a possible explanation why the relative gene expression varies somewhat with afuA. minC expression was at the same level in the AaMnR strain regardless of iron availability, and very close in iron-chelated medium, since these cells grew at a similar rate. The results from this experiment indicate that minC is iron regulated through the Fur protein. Thus, the Furta, the gel shift assays, and the RT-qPCR assays for gene expression all indicate that A. actinomycetemcomitans minC is Fur regulated. To our knowledge this is the first report of Fur regulation of the minC gene.
Conclusions
This study identified 14 putative Fur-regulated genes in A. actinomycetemcomitans. Many of these genes are known to be iron regulated in other bacterial species but one gene, minC, not previously described as being iron regulated, was found to be iron/Fur regulated in A. actinomycetemcomitans. A manganese-resistant fur mutant of A. actinomycetemcomitans was generated and shown to produce a non-functional Fur protein in our assays. The expressed protein might have retained some function; this will be the subject of future studies. The mutant strain was more sensitive to acid stress and expressed higher levels of minC than the wild-type strain.
The importance of these studies will be to improve our understanding of the nature of iron availability with regard to the pathogenesis of periodontal disease. In the long term, these studies will provide a firm foundation on which future investigations may be based that lead to the development of treatment to prevent or retard the progression of periodontal disease. It is possible that bacterial iron metabolism can be exploited as a new approach to antimicrobial chemotherapy. An intriguing aspect of this work is that, because iron is absolutely essential for bacteria, it could make the development of bacterial resistance very difficult for any organism deprived of iron. Chelation of iron in order to inhibit growth of invading bacteria could be developed into a therapy against periodontal infection. Another, even more intriguing possibility is that the identification of Fur-regulated proteins and their role in the pathogenesis of A. actinomycetemcomitans may facilitate the development of antibacterial drugs based on concepts such as delivering siderophore–antibiotic conjugates (Ghosh et al., 1996) with high affinity and specificity, through bacterial iron transport systems.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTS AND DISCUSSIONREFERENCES |
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Ang, S., Lee, C. Z., Peck, K., Sindici, M., Matrubutham, U., Gleeson, M. A. & Wang, J. T. (2001). Acid-induced gene expression in Helicobacter pylori: study in genomic scale by microarray. Infect Immun 69, 1679–1686.
Baehni, P. C., Tsai, C. C., McArthur, W. P., Hammond, B. F., Shenker, B. J. & Taichman, N. S. (1981). Leukotoxic activity in different strains of the bacterium Actinobacillus actinomycetemcomitans isolated from juvenile periodontitis in man. Arch Oral Biol 26, 671–676.[CrossRef][Medline]
Bagg, A. & Neilands, J. B. (1987). Ferric uptake regulation protein acts as a repressor, employing iron(II) as a cofactor to bind the operator of an iron transport operon in Escherichia coli. Biochemistry 26, 5471–5477.[CrossRef][Medline]
Bearden, S. W. & Perry, R. D. (1999). The Yfe system of Yersinia pestis transports iron and manganese and is required for full virulence of plague. Mol Microbiol 32, 403–414.[CrossRef][Medline]
Benson, H. P., LeVier, K. & Guerinot, M. L. (2004). A dominant-negative fur mutation in Bradyrhizobium japonicum. J Bacteriol 186, 1409–1414.
Berish, S. A., Subbarrao, S., Chen, C. V., Trees, D. L. & Morse, S. A. (1993). Identification and cloning of a fur homolog from Neisseria gonorrhoeae. Infect Immun 61, 4599–4606.
Beyer-Sehlmeyer, G., Kreikemeyer, B., Horster, A. & Podbielski, A. (2005). Analysis of the growth phase-associated transcriptome of Streptococcus pyogenes. Int J Med Microbiol 295, 161–177.[CrossRef][Medline]
Bijlsma, J. J., Waidner, B., Vliet, A. H. & 8 other authors (2002). The Helicobacter pylori homologue of the ferric uptake regulator is involved in acid resistance. Infect Immun 70, 606–611.
Brickman, T. J. & Armstrong, S. K. (1995). Bordetella pertussis fur gene restores iron repressibility of siderophore and protein expression to deregulated Bordetella bronchiseptica mutants. J Bacteriol 177, 268–270.
Bsat, N. & Helmann, J. D. (1999). Interaction of Bacillus subtilis Fur (ferric uptake repressor) with the dhb operator in vitro and in vivo. J Bacteriol 181, 4299–4307.
Christersson, L. A., Albini, B., Zambon, J. J., Wikesjo, U. M. & Genco, R. J. (1987). Tissue localization of Actinobacillus actinomycetemcomitans in human periodontitis. I. Light, immunofluorescence and electron microscopic studies. J Periodontol 58, 529–539.[Medline]
Clyne, M., Labigne, A. & Drumm, B. (1995). Helicobacter pylori requires an acidic environment to survive in the presence of urea. Infect Immun 63, 1669–1673.[Abstract]
de Boer, P. A., Crossley, R. E. & Rothfield, L. I. (1990). Central role for the Escherichia coli minC gene product in two different cell division-inhibition systems. Proc Natl Acad Sci U S A 87, 1129–1133.
de Boer, P. A., Crossley, R. E., Hand, A. R. & Rothfield, L. I. (1991). The MinD protein is a membrane ATPase required for the correct placement of the Escherichia coli division site. EMBO J 10, 4371–4380.[Medline]
de Lorenzo, V., Wee, S., Herrero, M. & Neilands, J. B. (1987). Operator sequences of the aerobactin operon of plasmid ColV-K30 binding the ferric uptake regulation (fur) repressor. J Bacteriol 169, 2624–2630.
Escolar, L., Perez-Martin, J. & de Lorenzo, V. (1999). Opening the iron box: transcriptional metalloregulation by the Fur protein. J Bacteriol 181, 6223–6229.
Fine, D. H., Velliyagounder, K., Furgang, D. & Kaplan, J. B. (2005). The Actinobacillus actinomycetemcomitans autotransporter adhesin Aae exhibits specificity for buccal epithelial cells from humans and old world primates. Infect Immun 73, 1947–1953.
Funahashi, T., Fujiwara, C., Okada, M., Miyoshi, S., Shinoda, S., Narimatsu, S. & Yamamoto, S. (2000). Characterization of Vibrio parahaemolyticus manganese-resistant mutants in reference to the function of the ferric uptake regulatory protein. Microbiol Immunol 44, 963–970.[Medline]
Ghosh, A., Ghosh, M., Niu, C., Malouin, F., Moellmann, U. & Miller, M. J. (1996). Iron transport-mediated drug delivery using mixed-ligand siderophore-beta-lactam conjugates. Chem Biol 3, 1011–1019.[CrossRef][Medline]
Grass, G., Franke, S., Taudte, N., Nies, D. H., Kucharski, L. M., Maguire, M. E. & Rensing, C. (2005). The metal permease ZupT from Escherichia coli is a transporter with a broad substrate spectrum. J Bacteriol 187, 1604–1611.
Hanahan, D. (1983). Studies on transformation of Escherichia coli with plasmids. J Mol Biol 5, 557–580.
Hantke, K. (1981). Regulation of ferric iron transport in Escherichia coli K12: isolation of a constitutive mutant. Mol Gen Genet 182, 228–292.
Hantke, K. (1987). Selection procedure for deregulated iron transport mutants (fur) in Escherichia coli K12: fur not only affects iron metabolism. Mol Gen Genet 210, 135–139.[CrossRef][Medline]
Haraszthy, V. I., Lally, E. T., Haraszthy, G. G. & Zambon, J. J. (2002). Molecular cloning of the fur gene from Actinobacillus actinomycetemcomitans. Infect Immun 70, 3170–3179.
Hayashida, H., Poulsen, K. & Kilian, M. (2002). Differences in iron acquisition from human haemoglobin among strains of Actinobacillus actinomycetemcomitans. Microbiology 148, 3993–4001.
Helgeland, K. & Nordby, O. (1993). Cell cycle-specific growth inhibitory effect on human gingival fibroblasts of a toxin isolated from the culture medium of Actinobacillus actinomycetemcomitans. J Periodont Res 28, 161–165.[CrossRef][Medline]
Hoegy, F., Celia, H., Mislin, G. L., Vincent, M., Gallay, J. & Schalk, I. J. (2005). Binding of iron-free siderophore, a common feature of siderophore outer membrane transporters of Escherichia coli and Pseudomonas aeruginosa. J Biol Chem 280, 20222–20230.
Huesca, M., Goodwin, A., Bhagwansingh, A., Hoffman, P. & Lingwood, C. A. (1998). Characterization of an acidic-pH-inducible stress protein (Hsp70), a putative sulfatide binding adhesin, from Helicobacter pylori. Infect Immun 66, 4061–4067.
Kaplan, A. H., Weber, D. J., Oddone, E. Z. & Perfect, J. R. (1989). Infection due to Actinobacillus actinomycetemcomitans: 15 cases and review. Rev Infect Dis 11, 46–63.[Medline]
Kolade, O. O., Bellini, P., Wexler, M., Johnston, A. W. B., Grossmann, J. G. & Hemmings, A. M. (2001). Structural studies of the Fur protein from Rhizobium leguminosarum. Biochem Soc Trans 30, 771–774.
Komatsuzawa, H., Asakawa, R., Kawai, T., Ochiai, K., Fujiwara, T., Taubman, M. A., Ohara, M., Kurihara, H. & Sugai, M. (2002). Identification of six major outer membrane proteins from Actinobacillus actinomycetemcomitans. Gene 288, 195–201.[CrossRef][Medline]
Lam, M. S., Litwin, C. M., Carroll, P. A. & Calderwood, B. (1994). Vibrio cholerae fur mutations associated with loss of repressor activity: implications for the structural–functional relationships of fur. J Bacteriol 176, 5108–5115.
Litwin, C. M. & Calderwood, S. B. (1993). Role of iron in the regulation of virulence genes. Clin Microbiol Rev 6, 137–149.
Litwin, C. M. & Calderwood, S. B. (1994). Analysis of the complexity of gene regulation by fur in Vibrio cholerae. J Bacteriol 176, 240–248.
Loosmore, S. M., Yang, Y. P., Coleman, D. C., Shortreed, J. M., England, D. M., Harkness, R. E., Chong, P. S. & Klein, M. H. (1996). Cloning and expression of the Haemophilus influenzae transferrin receptor genes. Mol Microbiol 19, 575–586.[CrossRef][Medline]
McGowan, C. C., Necheva, A., Thompson, S. A., Cover, T. L. & Blaser, M. J. (1998). Acid-induced expression of an LPS-associated gene in Helicobacter pylori. Mol Microbiol 30, 19–31.[CrossRef][Medline]
Merrell, D. S., Hava, D. L. & Camilli, A. (2002). Identification of novel factors involved in colonization and acid tolerance of Vibrio cholerae. Mol Microbiol 43, 1471–1491.[CrossRef][Medline]
Merrell, D. S., Goodrich, M. L., Otto, G., Tompkins, L. S. & Falkow, S. (2003). pH-regulated gene expression of the gastric pathogen Helicobacter pylori. Infect Immun 71, 3529–3539.
Messing, J. (1993). M13 cloning vehicles. Their contribution to DNA sequencing. Methods Mol Biol 23, 9–22.[Medline]
Meyer, D. H. & Fives-Taylor, P. M. (1993). Evidence that extracellular components function in adherence of Actinobacillus actinomycetemcomitans to epithelial cells. Infect Immun 61, 4933–4936.
Miller, J. H. (1972). Experiments in Molecular Genetics. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory.
Mintz, K. P. & Fives-Taylor, P. M. (1994). Identification of an immunoglobulin Fc receptor of Actinobacillus actinomycetemcomitans. Infect Immun 62, 4500–4505.
Mintz, K. P. & Fives-Taylor, P. M. (1999). Binding of the periodontal pathogen Actinobacillus actinomycetemcomitans to extracellular matrix proteins. Oral Microbiol Immunol 14, 109–116.[CrossRef][Medline]
Mombelli, A., Gmur, R., Gobbi, C. & Lang, N. P. (1994). Actinobacillus actinomycetemcomitans in adult periodontitis. II. Characterization of isolated strains and effect of mechanical periodontal treatment. J Periodontol 65, 827–834.[Medline]
Moran, A. P., Knirel, Y. A., Senchenkova, S. N., Widmalm, G., Hynes, S. O. & Jansson, P. E. (2002). Phenotypic variation in molecular mimicry between Helicobacter pylori lipopolysaccharides and human gastric epithelial cell surface glycoforms. Acid-induced phase variation in Lewisx and Lewisy expression by H. pylori lipopolysaccharides. J Biol Chem 277, 5785–5795.
Ochsner, U. A., Vasil, A. I. & Vasil, M. L. (1995). Role of the ferric uptake regulator of Pseudomonas aeruginosa in the regulation of siderophores and exotoxin A expression: purification and activity on iron-regulated promoters. J Bacteriol 177, 7194–7201.
Overholt, B. F. (1966). Actinobacillus actinomycetemcomitans endocarditis. Arch Intern Med 117, 99–102.[CrossRef][Medline]
Palyada, K., Threadgill, D. & Stintzi, A. (2004). Iron acquisition and regulation in Campylobacter jejuni. J Bacteriol 186, 4714–4729.
Pancholi, V. (2001). Multifunctional alpha-enolase: its role in diseases. Cell Mol Life Sci 58, 902–920.[CrossRef][Medline]
Pancholi, V. & Fischetti, V. A. (1998). Alpha-enolase, a novel strong plasmin(ogen) binding protein on the surface of pathogenic streptococci. J Biol Chem 273, 14503–14515.
Prince, R. W., Cox, C. D. & Vasil, M. L. (1993). Coordinate regulation of siderophore and exotoxin A production: molecular cloning and sequencing of the Pseudomonas aeruginosa fur gene. J Bacteriol 175, 2589–2598.
Ratledge, C. & Dover, L. G. (2000). Iron metabolism in pathogenic bacteria. Annu Rev Microbiol 54, 881–941.[CrossRef][Medline]
Rhodes, E. R., Tomaras, A. P., McGillivary, G., Connerly, P. L. & Actis, L. A. (2005). Genetic and functional analyses of the Actinobacillus actinomycetemcomitans AfeABCD siderophore-independent iron acquisition. Infect Immun 73, 3758–3763.
Roe, B. A., Najar, F. Z., Gillaspy, A., Clifton, S., Ducey, T., Lewis, L. & Dyer, D. W. (1999). Actinobacillus Genome Sequencing Project, University of Oklahoma. http://www.genome.ou.edu/act.html
Sambrook, J., Fritsch, E. F. & Maniatis, T. (1989). Molecular Cloning: a Laboratory Manual, 2nd edn. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory.
Sanger, F., Nicklen, S. & Coulson, A. R. (1977). DNA sequencing with chain-terminating inhibitors. Proc Natl Acad Sci U S A 74, 5463–5467.
Schaffer, S., Hantke, K. & Braun, V. (1985). Nucleotide sequence of the iron regulatory gene fur. Mol Gen Genet 200, 110–113.[CrossRef][Medline]
Stojiljkovic, I., Baumler, A. J. & Hantke, K. (1994). fur regulon in gram-negative bacteria. Identification and characterization of new iron-regulated Escherichia coli genes by a fur titration assay. J Mol Biol 236, 531–545.[CrossRef][Medline]
Thomas, C. E. & Sparling, P. F. (1996). Isolation and analysis of a fur mutant of Neisseria gonorrhoeae. J Bacteriol 178, 4224–4232.
Tolmasky, M. E., Wertheimer, A. M., Actis, L. A. & Crosa, J. H. (1994). Characterization of the Vibrio anguillarum fur gene: role in regulation of expression of the FatA outer membrane protein and catechols. J Bacteriol 176, 213–220.
Touati, D., Jacques, M., Tardat, B., Bouchard, L. & Despied, S. (1995). Lethal oxidative damage and mutagenesis are generated by iron in fur mutants of Escherichia coli: protective role of superoxide dismutase. J Bacteriol 177, 2305–2314.
Townsend, T. R. & Gillenwater, J. Y. (1969). Urinary tract infection due to Actinobacillus actinomycetemcomitans. JAMA 210, 558.[CrossRef][Medline]
Tsolis, R. M., Bäumler, A. J., Stojiljkovic, I. & Heffron, F. (1995). Fur regulon of Salmonella typhimirium: identification of new iron-regulated genes. J Bacteriol 177, 4628–4637.
van Vliet, A. H., Rock, J. D., Madeleine, L. N. & Ketley, J. M. (2000). The iron-responsive regulator Fur of Campylobacter jejuni is expressed from two separate promoters. FEMS Microbiol Lett 188, 115–118.[Medline]
Willemsen, P. T., Vulto, I., Boxem, M. & de Graaff, J. (1997). Characterization of a periplasmic protein involved in iron utilization of Actinobacillus actinomycetemcomitans. J Bacteriol 179, 4949–4952.
Winer, J., Jung, C. K. S., Shackel, I. & Williams, P. M. (1999). Development and validation of real-time quantitative reverse transcriptase-polymerase chain reaction for monitoring gene expression in cardiac myocytes in vitro. Anal Biochem 270, 41–49.[CrossRef][Medline]
Wurgler, S. M. & Richardson, C. C. (1990). Structure and regulation of the gene for dGTP triphosphohydrolase from Escherichia coli. Proc Natl Acad Sci U S A 87, 2740–2744.
Zambon, J. J. (1994). Actinobacillus actinomycetemcomitans in adult periodontitis. J Periodontol 65, 892–893.[Medline]
Received 23 July 2005;
revised 12 October 2005;
accepted 17 November 2005.
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) to that of Aawt cells (
) under iron-rich conditions: AaMnR cells grew more slowly than Aawt cells under these conditions. Under iron-chelated conditions the growth of AaMnR cells (
) was very similar to that of Aawt cells (
) type growth were very similar. The results are means of three independent growth experiments done in triplicate.
) after 24 h (stationary phase) in broth adjusted to pH from 6·1 to 7·2 in increments of 0·1 unit. The parent strain was able to achieve limited growth at pH 6·1 but the fur mutant was not able to grow at that pH. Growth was restored at pH 6·9 for the AaMnR strain while Aawt grew at 6·5 pH at its normal rate.
