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Department of Microbiology and Immunology, University of Western Ontario, London, Ontario, Canada
Received 9 September 2003/ Accepted 9 January 2004
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ABSTRACT |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Several virulence factors contribute to the stringent host and tissue specificities exhibited by H. pylori (37). Among them, urease helps neutralize the acidic pH surrounding the bacteria and allows their survival in the gastric environment (21, 45). In addition, the spiral shape and unipolar flagella of H. pylori confer on the bacterium a corkscrew motion that enhances motility in the viscous gastric mucus (32, 33, 63) and is essential for host colonization (22, 23, 35). Lipopolysaccharide (LPS) is also important for the virulence of H. pylori since strains lacking the O antigen are significantly impaired in their capacity to colonize the murine stomach (40). The H. pylori O antigen is composed of N-acetyl-D-glucosamine, L-fucose, and D-galactose (4, 5, 46), which form structural motifs that are identical to human blood group antigens Lewis X, Y, and b (4, 5, 46-48). This host mimicry might allow the bacteria to evade human immune defenses and to establish long-term host colonization (54, 61).
This study focuses on two genes of unknown biological function in H. pylori: flaA1 (HP0840) and wbpB (HP0679). They exhibit significant homologies to LPS (9), capsule (8, 57, 58), and/or flagellar biosynthetic genes found in medically relevant bacteria. For example, FlaA1 shows 52% homology to the C-terminal half of WbpM, which is essential for LPS synthesis in Pseudomonas aeruginosa (9). The sequence homologies result in functional equivalence between these proteins, since flaA1 can support O-antigen biosynthesis in a wbpM mutant (15). Additional homologues identified in Campylobacter jejuni (PglF and CJ1293) (25, 64) and Caulobacter crescentus (FlmA) (38) are involved in protein glycosylation and/or influence flagellum production. In contrast to FlaA1, which seems widely distributed in the bacterial world, WbpB has only two homologues in bacterial genomes. One (WbpBPa, 63% homology) is found in the LPS biosynthetic cluster of P. aeruginosa serotype O5 (9), and the other (WlbA, 52% homology, N-terminal half only) is found in the LPS biosynthetic cluster of Bordetella pertussis (1). The homologies described above strongly suggest that flaA1 and wbpB might also be involved in LPS and flagellum biosynthesis in H. pylori. Interestingly, in bacteria where a homologue for each gene is found, these homologues are found together within a cluster of genes that are dedicated to the same biological function, suggesting a potential functional link for the H. pylori genes despite their presence in distinct areas of the chromosome.
Both genes encode sugar-nucleotide-modifying enzymes. FlaA1 is a UDP-GlcNAc C-6 dehydratase/C-4 reductase (15, 16), and WbpB is predicted to be an oxidoreductase. Consistent with the potential functional link between flaA1 and wbpB mentioned above, WbpB might be involved in the reduction of the UDP-4-keto-6-deoxy-GlcNAc intermediate generated by FlaA1. Interestingly, neither the 4-keto intermediate nor its reduced derivative are present in the LPS of H. pylori (4, 5, 46). In addition, all genes involved in the biosynthesis of the precursors necessary for LPS assembly have been identified (30, 36, 44) and are distinct from flaA1 and wbpB. Consequently, although it is expected that flaA1 and wbpB might affect LPS synthesis based on sequence homologies, such an effect is not anticipated to occur directly via production of LPS sugar precursors.
The sequence homologies described above also suggested that flaA1 and wbpB might be involved in protein glycosylation. This is relevant since flagellin glycosylation has been detected in H. pylori (34, 60) and it is possible that additional proteins—other than the flagellins—are glycosylated, as is the case in the closely related C. jejuni (64, 72).
Using a gene disruption strategy, we investigated the biological function of flaA1 and wbpB in H. pylori. In light of their homologies to LPS biosynthetic and flagellum modification genes found in other bacteria, the study was focused on the role of both genes on LPS biosynthesis and outer membrane barrier function, as well as on their role in flagellum synthesis and function.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Cloning experiments were performed with Escherichia coli DH5
unless stated otherwise, using 30 µg of kanamycin per ml, 34 µg of chloramphenicol per ml, or 100 µg of ampicillin per ml when necessary. Each construct was checked by restriction analysis and sequencing. All kits were used as specified by their manufacturer. DNA sequencing was performed at the Robarts Institute Sequencing Facility (London, Canada).
Cloning and sequencing of the flaA1 and wbpB genes from strains SS1 and NCTC 11637. wbpB and flaA1 were amplified from chromosomal DNA using Pwo (Roche) DNA polymerase and primers HPWB5 (GCTCTCCATGGGTATGCTTTTTGCGATGATTG) and HPWB3 (AAGCAGGATCCTCAAGCCAATTTGACAGACG) for wbpB and Flatop (ACTGTACATGTCAATGCCAAATCATCAAAAC) and Flabot (AAGCTGGATCCTCATAATAATTTCAACAAA) for flaA1. Both PCR products were cloned in Topo-PCR2.1 (Invitrogen) using Top10F' cells. The constructs were sequenced on both strands using M13 forward and reverse primers.
Cloning and sequencing of the flagellin genes, flaA and flaB, from strain NCTC 11637. flaA and flaB were amplified from chromosomal DNA using primers based on the sequences of HP0601 and HP0115 from strain 26695. Note that flaA (HP0601, encoding the flagellin) is distinct from flaA1 (HP0840, encoding the enzyme under study). The primers were HPFlaAP3 (GCTCTCCATGGCTTTTCAGGTCAATAC) and HPFlaAP5 (AAGAAGATCTCCTAAGTTAAAAGCCTTAAG) for flaA and HP0115P1 (GCTCTCCATGGGCATGAGTFTTAGGATAAATAC) and HP0115P2 (AAGAGGATCCTTATTGTAAAAGCCTTAAGA) for flaB. The PCR was performed using an Expand long-range template (Roche) with annealing at 46°C. Both PCR products were cloned in the pET23a derivative (13, 49) with an N-terminal histidine tag using NcoI and BglII for flaA and NcoI and BamHI for flaB. The constructs were sequenced using T7 promoter primer and primers HPFlagAP3 (GTGAATGATGTAACTTTAGAG) and HPFlagBP3 (CTTATAATGTCATGGCTACC).
Production of anti-flagellin A antibody. Flagellin A was overexpressed from the pET23 construct in BL21(DE3)pLys cells with induction by 0.5 mM isopropyl-ß-D-thiogalactopyranoside (IPTG) at 37°C. The protein was purified by metal chelation as described previously (13) in the presence of 6 M guanidine-HCl. The protein was dialyzed in 50 mM ammonium bicarbonate in the presence of 1% sodium dodecyl sulfate (SDS), subjected to microelution from SDS-polyacrylamide gel electrophoresis (PAGE) gels (Bio-Rad Electro-Eluter), and redialyzed in the same buffer. The dialyzate was lyophilized, resuspended in 1.7% saline, and used to immunize rabbits after 1:1 (vol/vol) dilution with incomplete Freund's adjuvant. The immunization schedule and procedure for adsorption of the collected serum against an E. coli BL21(DE3)pLys extract were as reported previously (14).
To further enhance the specificity of the serum toward flagellins, the serum was passed through a Blue Sepharose 6 Fast Flow column (Pharmacia Biotech) and purified onto a flagellin A affinity column that was made by coupling purified overexpressed flagellin A to CNBr-activated Sepharose (Pharmacia Biotech) in the presence of guanidine-HCl. After being washed with 1 column volume (CV) of 150 mM glycine (pH 8.9)-350 mM NaCl and after acid shock with 1 CV of 100 mM glycine (pH 3.2), flagellin-specific antibodies were eluted with 1 CV of 1 M ammonium hydroxide and immediately neutralized with 0.09 CV of 0.2 M acetic acid. The recovered antibodies were further adsorbed against a lysate from a H. pylori FlaA knockout mutant (see below) as described previously (14).
Preparation of the knockout and complementation constructs. (i) flaA1 knockout construct. flaA1 (HP0840) and its potential promoter were PCR amplified from genomic DNA using Taq DNA polymerase (Invitrogen) and primers FlaTop2 (GCGAGCGCGAATCTTTAT) and FlaA1Top (ATAGAACCGCTCACGAGC). The PCR product was cloned into Topo-PCR2.1 to generate TopoFla and was subsequently subcloned into the BamHI and ApaI sites of pBluescript II SK. The construct was digested with NcoI and blunted using T4 DNA polymerase. The kanamycin resistance cassette and its promoter were cut out from pHel3 (27) using SmaI and ligated into the blunted flaA1-pBluescript fragment to generate the flaA1 knockout construct.
(ii) wbpB knockout construct. wbpB (HP0679) and its potential promoter were PCR amplified from genomic DNA using primers WbpBup (AACAGAGCCCACGAACGA) and WbpBdown (ATCACGCTTGCGATTGGC) and Taq DNA polymerase. The PCR product was cloned into Topo-PCR2.1 to generate TopoWbpB and subcloned into the EcoRI site of pBluescript II SK to generate pBSWbpB. The construct was digested with NsiI and blunted using T4 DNA polymerase. The kanamycin resistance cassette was cut from pHel3 (27) using SmaI and ligated into the blunted pBSWbpB fragment to generate the wbpB knockout construct.
(iii) flaA knockout construct. Inverse PCR amplification of the flaA/pET23 construct was performed with primers HPFlagAP3 (GTGAATGATGTAACTTTAGAG) and HPFlagAP4 (GAACGATGTCAGATTGAATC) and Expand long-range template polymerase (Roche). The PCR product was blunted and dephosphorylated before being ligated to the SmaI-extracted kanamycin cassette as described above.
(iv) Complementation constructs. The EcoRI fragment containing either gene with its promoter was cut out from TopoFla or TopoWbpB, blunted with T4 DNA polymerase, and cloned into the EcoRV-cut pHel2 shuttle vector (27) to generate the complementation constructs pHel2-FlaA1 and pHel2-WbpB.
Southern blotting. Southern blotting was performed using the digoxigenin-labeling method with detection with anti-digoxigenin-alkaline phosphatase-CSPD (disodium 3-(4-methoxyspiro{1,2-dioxetane-3,2'-(5'-chloro)tricyclo[3.3.1.13,7]decan}-4-yl)phenyl phosphate) substrate as specified by the manufacturer (Roche). Chromosomal DNA was extracted from H. pylori using DNAzol (Invitrogen) and cut with HaeII/HindIII (flaA1 mutant) or MaeIII (wbpB mutant).
Preparation of knockout mutants and complemented clones. The knockout mutants were generated by electroporation-mediated allelic exchange (63) with modifications described by McGee et al. (43) and selection on kanamycin. Potential transformants were analyzed for gene integration by PCR and Southern blotting. Complementation constructs were introduced into the mutants by electroporation, with selection onto kanamycin and chloramphenicol. The plasmids were extracted from the complemented strains and subjected to restriction analysis to ensure their integrity. Transformants harboring intact plasmids were selected for further studies.
LPS analysis. LPS prepared as described previously (28) was analyzed on SDS-PAGE (15% polyacrylamide) or 10 to 20% Tricine gradient (Novex) gels. Detection was performed by silver staining (24) or Western blotting with anti-Lewis Y (Calbiochem) or anti-lipid A monoclonal antibodies (19).
Preparation of soluble cell extracts. Soluble extracts were obtained by lysing H. pylori cells harvested from one BHI-YE plate in 100 µl of breaking buffer (20 mM sodium phosphate [pH 7.5], 1 mM EDTA) with acid-washed glass beads (Sigma G-4649). After being vortexed three times for 30 s each, the cells were pelletted for 10 min at 12,000 x g at room temperature and the supernatant was used for SDS-PAGE analysis.
Protein deglycosylation. Enzymatic deglycosylation of flagellum preparations obtained by glycine extraction (50) was performed under denaturing conditions using five different enzymes as provided in the Calbiochem glycoprotein deglycosylation kit (no. 362280).
MS identification of proteins. Protein bands cut out from Coomassie-stained gels were subjected to in-gel trypsinolysis. The peptides were analyzed by liquid chromatography mass spectrometry (MS) (Q-TOF2) and MALDI-MS at the Dr. Don Rix Protein Identification Facility of the University of Western Ontario.
RT-PCR. RNA extracted using the RNeasy kit (Qiagen) was treated with RNase-free DNase I (Invitrogen) and subjected to reverse transcription (RT) using Superscript II RNase H- reverse transcriptase (Invitrogen) and random hexanucleotide primers (Roche). PCR (15 cycles) was subsequently performed with Taq DNA polymerase at an annealing temperature of 52°C and elongation time of 45 s, using gene-specific primers. The primers were Fla363 (GCTATCAGTCAGGTTATCGC) and Fla550 (ACGGCACCACGCTCCCAC) to amplify a 187-bp fragment of flaA1, HPWB8 (CAGAACACATGGGAGTAGC) and HPWB7 (GCCGTCCGAGCGCCAATTTGACAGACGC) to amplify a 171-bp fragment of wbpB, and HP1045P1 (GTCATTATCTATATGCCCAT) and HP1045P2 (CTGGCTTGAGCATGTAAGG) to amplify a 200-bp fragment from HP1045. The PCR products were analyzed on 2% agarose gels with ethidium bromide staining.
SDS, bile salts, and novobiocin sensitivity assays. Harvested bacteria were diluted to an optical density at 600 nm (OD600) of 1 in BHI-YE. A 30-µl volume of cell suspension was added to 175 µl of BHI-YE supplemented with the appropriate antibiotics and/or detergents, with concentrations up to 15 µg/ml for novobiocin, 0.02% for SDS, and 0.18 mg/ml for bile salts (50% sodium cholate, 50% sodium deoxycholate (Sigma) (29). The assays were performed three times with independent cultures in 96-well plates incubated for 4 to 5 days with agitation at 37°C under microaerophilic conditions. On each plate, each strain was tested in duplicate. A representative example of the SDS-sensitivity experiment is provided in Fig. 6.
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Motility assays. Bacteria were harvested from a 2-day-old plate into 400 µl of BHI-YE or from a 1-ml overnight liquid culture, diluted 1:10 in BHI-YE saturated with 10% CO2-85% N2-5% O2, and allowed to grow for 7.5 h. The cultures were then diluted to an OD600 of 0.3 or 0.9 and were used to inoculate motility plates (0.4% agar in BHI-YE) by stabbing. Each dilution was spotted in triplicate, and dilutions were made in triplicate for each strain. The growth of the swimming halo was monitored after 4 days of incubation at 37°C. The remaining diluted culture was used to inoculate regular BHI-YE plates to estimate bacterial viability by colony counting and microscopic observation.
Suspension-clearing assays. Liquid cultures (1 ml) in BHI-YE saturated with 10% CO2-85% N2-5% O2 were prepared by inoculation from 2-day-old plates. After overnight growth, the cultures were diluted 1:10 in 2 ml of BHI and grown again for 18 to 24 h. The cultures were then diluted to an OD600 of 1 (total volume, 2 ml) in BHI and left to sit at room temperature. The rate of suspension clearance was measured over 8 h by carefully removing 70 µl from the top of the suspension and measuring the OD600 using a microcell (path length, 1 cm). Two independent experiments using two dilutions of each strain were performed.
Microscopic observations. Cells harvested from plates or liquid cultures were examined as wet mounts under a phase-contrast microscope (Zeiss; oil immersion, x400 magnification). For electron microscopy, cells were harvested from 1- to 2-day-old plates and resuspended in 1% glutaraldehyde in 20 mM HEPES buffer (pH 7). They were analyzed in the negative staining mode using 1% uranyl acetate.
Urease activity assay. A 10-µl volume of a bacterial suspension at an OD600 of 0.4 in saline was mixed with 100 µl of 0.33 M urea-0.001% phenol red in 0.005 M sodium phosphate-0.15 M NaCl (pH 6.7) (11). Urease activity was assayed by monitoring the OD565 over time in a 96-well plate. All experiments were done in triplicate. The data were normalized to the total amount of proteins present in the sample (Bio-Rad assay).
Growth curves. Bacterial suspensions (1 ml at an OD600 of 0.055) in BHI-YE were grown in 24-well plates with agitation under microaerophilic conditions, and the OD600 was monitored over 70 h. Data were recorded from four different plates with two repeats per strain on each plate. The data presented (see Fig. 2A) are the average of all the readings.
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Nucleotide sequence accession numbers. The GenBank accession numbers for the genes in this study are as follows: flaA1 (NCTC 11637), AY319294; flaA1 (SS1), AY319295; wbpB (NCTC 11637), AY319297; wbpB (SS), AY319296; flaA (NCTC 11637), AY319298; and flaB (NCTC 11637), AY319299.
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RESULTS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Like the wild type, both mutants were able to resist acid shock in the absence of urea at pH 
3 (Fig. 3A to C, bars a). The addition of urea allowed resistance to acid shock of all strains at pH 2 (bars b to e). This indicated that the limited urease activity of the flaA1 mutant was sufficient to support viability at pH 2. At pH 7, addition of urea became deleterious. In the flaA1 mutant, this toxic effect inherent to the chaotropic nature of urea appeared at lower urea concentrations (15 mM) (Fig. 3B, bars d and e) than for the wild type and for all pHs of >2. This was consistent with the lower levels of urease activity shown by this mutant (Fig. 2B).
Like wild-type bacteria, both mutants resisted long-term acid exposure in the absence of urea (Fig. 3D to F, bars a). The slight decrease in viability observed for all strains at pH 5 could be eliminated by the addition of 10 mM urea (bars c), indicating again that urease activity supports acid resistance under these conditions too.
The flaA1 and wbpB knockout mutants exhibit altered LPS profiles. SDS-PAGE analysis of wild-type and mutant LPS revealed that the mutants had lost most of their O antigen (Fig. 4A). The wbpB mutant produced slightly more Lewis Y-containing O antigen than did the flaA1 mutant. Also, the O chains present in the wbpB mutant appeared slightly shifted compared with the wild-type bands (Fig. 4B), suggesting that the core to which the O antigen was attached might have a slightly different structure. When the core LPS was analyzed on higher-resolution 10 to 20% gradient Tricine gels, the wild-type strain exhibited two well resolved bands, bands 1 and 2 (Fig. 4C). In contrast, in both mutants, the fast-migrating band (band 1) was missing and the slower-migrating band (band 2) was not affected. In addition, a third band (band 3) migrating slightly slower than band 2 appeared in the wbpB mutant only. The presence of band 3 was not dependent on the amount of sample loaded on the gel.
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Reintroducing the functional gene in trans partly restored the production of the O antigen in each mutant (Fig. 5), indicating that the loss of O-antigen production observed in each mutant was gene specific and not due to polarity effects. The copy number and poor stability of the complementation plasmid in strain NCTC 11637 might explain why complementation was not complete. In the flaA1-complemented flaA1 mutant, the O antigen produced was composed of Lewis Y-containing species, as in the wild type (white and hatched bars of similar size). Interestingly, in the wbpB-complemented wbpB mutant, the amount of Lewis Y-containing O antigen did not increase (hatched bar) but its proportion of the total O antigen (detected with anti-lipid A [white bar]) did increase, so that all O antigen produced was of the Lewis Y type, as in the wild type. Hence, qualitatively, reintroducing the functional wbpB gene into the wbpB mutant also restored a wild-type like O-antigen production made exclusively of Lewis Y O antigens. The identity of the Lewis Y-reacting band that appears below the O antigen in the flaA1::kan + wbpB and flaA1::kan lanes (Fig. 5A) is unknown, but it does not appear to be LPS related since it did not react with anti-lipid A antibody.
Disruption of flaA1 or wbpB affects the barrier properties of the outer membrane. Defects in LPS synthesis often correlate with higher sensitivity of bacteria to killing by serum, detergents (SDS or bile salts) (29), or hydrophobic antibiotics (novobiocin) (67) and with decreased virulence properties in animal models (56, 70). Hence, the sensitivity of the flaA1 and wbpB mutants to each of these compounds was investigated.
The extreme sensitivity to serum of the NCTC 11637 strain used for these studies (7) prevented us from detecting any significant differences between the wild type and mutants. However, a significantly higher sensitivity of the mutants than of the wild type to SDS (Fig. 6A) and a slight increase in sensitivity to novobiocin and bile salt were observed (data not shown). These results indicate the higher accessibility of the outer membrane to antibiotic or detergents and its reduced stability, which is consistent with decreased O-antigen production.
Providing the functional gene in trans restored wild-type-like SDS sensitivity in each mutant (Fig. 6B and C), demonstrating the gene specificity of the increased SDS sensitivity phenotype.
flaA1 can cross-complement the wbpB inactivation. We investigated whether the bifunctional dehydratase/reductase FlaA1 could complement the disruption of the putative reductase activity of WbpB by monitoring both the LPS production by and SDS sensitivity of cross-complemented mutants. Providing a functional copy of flaA1 in trans in the wbpB mutant did restore wild-type-like SDS sensitivity to the wbpB mutant (Fig. 6C). This correlated with an increased production of Lewis Y-containing O antigen in this complemented mutant (Fig. 5, hatched bars). This indicated that FlaA1 and WbpB have a redundant reductase activity and might be involved in the same biochemical pathway. However, providing wbpB in the flaA1 mutant did not restore wild-type levels of SDS sensitivity (Fig. 6B) or O-antigen production (Fig. 5) to the flaA1 mutant, indicating that the complementation effect observed previously was gene specific. This is consistent with the fact that WbpB is predicted to be monofunctional (reductase only) and cannot complement the lack of dehydratase function of the flaA1 mutant.
The flaA1 and wbpB disruptions abrogate flagellum-mediated motility. On examination by phase-contrast microscopy, the wild-type bacteria were motile and exhibited corkscrew movements. In contrast, the flaA1 and wbpB mutants only oscillated and showed no corkscrew movement.
Motility assays performed with bacteria in soft agar showed a clear reduction of motility on disruption of the flaA1 and wbpB genes (Table 2). The results were not dependent on the length of incubation or on the density or nature (from broth or plate) of the inoculum.
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First, we demonstrated that strain NCTC 11637 produces two flagellins, FlaA and FlaB, which were >99.5% identical to flagellins found in strain 26695 and 57.2% identical to one another. NCTC 11637 FlaA was 53.3 kDa in size, and FlaB was only slightly larger (54.0 kDa).
Second, we demonstrated that both flagellins could be readily detected by Western immunoblotting using a polyclonal antiserum raised in rabbits against overexpressed and purified NCTC 11637 FlaA flagellin. This was consistent with their high levels of similarity at the protein level. Note that in the gel system used, the two flagellins were not resolved from one another.
Third, the anti-FlaA antibody was used to investigate flagellin production in wild-type and mutant H. pylori strains. Figure 9 shows that even after adsorption against an E. coli lysate, the anti-FlaA antibody detected several proteins in wild-type H. pylori soluble extracts (bands a to d). Band c (53 kDa) had the strongest reactivity with the anti-FlaA antibody and migrated to the same position as pure overexpressed flagellins (data not shown), indicating that it corresponds to the flagellin(s) FlaA and/or FlaB. Band c was not affected by the flaA1 or wbpB disruption, indicating that both mutants still produced flagellin(s), including the aflagelatted flaA1 mutant. Hence, the data suggest that the export and/or assembly of the flagellins is impaired in the flaA1 mutant.
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MS analysis indicated that band d contained multiple proteins unrelated to flagellins. Since this band was not affected by the flaA1 or wbpB disruptions, its analysis was not pursued further.
Finally, the production of band b (58 kDa) was abrogated in the flaA1 mutant only, and the effect was gene specific since band b reappeared in the flaA1-complemented flaA1 mutant (Fig. 9). Considering that several strains of H. pylori harbour glycosylated flagellins (34, 60), the possibility that band b might correspond to posttranslationally modified flagellins was investigated by MS analysis. However, contamination of band b with catalase (58.6 kDa) that was present both in the wild type and in the flaA1 mutant (a negative control that does not contain band b) prevented us from demonstrating the presence of flagellins within band b. The experiments below aim at clarifying this question by refining the purification and detection of the flagellins and by investigating the glycosylation status of the flagellins in the band b-producing wild-type and wbpB mutant strains.
The flagellins of wild-type H. pylori strain NCTC 11637 are not glycosylated. To examine the glycosylation status of the NCTC 11637 flagellins, flagellins obtained by glycine extraction were analyzed by Western immunoblotting as well as Coomassie and Ponceau staining (Fig. 10). Three major bands (bands A to C) in the size range of interest were detected by Coomassie (Fig. 10C) or Ponceau (Fig. 10A) staining. As described above (Fig. 9), band A is thought to be UreB. A significant reduction in reactivity of the serum with band A was obtained using serum that had been purified by affinity on a flagellin A column and adsorbed against a lysate of a urease-producing H. pylori flagellin A mutant (compare the intensity of band B in Fig. 10B and D).
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Enzymatic deglycosylation using a combination of five different enzymes had no effect on the migration or amount of band C, indicating that it corresponds to nonglycosylated flagellins. C. jejuni glycosylated flagellines prepared and treated under the same conditions were included as a positive control and showed a significant shift in size after deglycosylation. Band B could be readily deglycosylated enzymatically (Fig. 10C). However, since it did not react with the anti-FlaA antibody and did not comigrate with band C after deglycosylation, it does not correspond to glycosylated flagellins. Note that no equivalent of the anti-FlaA-reacting band b from Fig. 9 was observed in glycine extracts, indicating that the glycine extraction method eliminated cross-reacting bands that were not flagellum related.
Overall, this analysis excludes the existence of any glycosylated flagellins in the wild-type strain and suggests that the wbpB mutant produces wild-type-like flagellins. However, these data did not determine whether the lack of motility of the wbpB mutant could be due to a lack of production of one of the two flagellins since FlaA and FlaB could not be resolved on these gels, and even after adsorption against a flaA knockout mutant, the anti-FlaA antibody still detected both flagellins without discrimination (Fig. 10B). Attempts at refining the purity of the flagellins by ultracentrifugation/acid dissociation and affinity chromatography to answer this question by direct MS analysis were unsuccessful due to the precipitation and therefore the loss of the flagellins.
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DISCUSSION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The two genes are distant from one another on the H. pylori chromosome (2, 65). The similarity of the phenotypes observed after disruption of either gene suggested that they belonged to the same functional pathway despite the absence of genetic linkage. Indeed, each gene affects LPS synthesis, outer membrane barrier function, and flagellum-mediated motility. However, subtle differences in the phenotypes of each mutant could be observed, and the phenotypes were proven to be gene specific by complementation experiments.
The functional link between the two genes is consistent with the fact that in the few bacteria that harbor a homologue for each gene, both genes were found within a single operon and participated in the same biological function (1, 9). This is also consistent with their biochemical activities, which might be partly complementary. Indeed, the unidirectional complementation of the disruption of wbpB by flaA1 supports the hypothesis that this functional complementation involves the common reductase activity associated with each enzyme. While this awaits further biochemical evidence, the reason for such redundancy of the reductase activity is currently not clear. The chromosomally encoded copy of flaA1 only partially complemented the wbpB mutation (Fig. 5 and 6C), so that the phenotype observed in the wbpB mutant was intermediate between those of the wild type and the flaA1 mutant (with the presence of flagella and higher levels of residual O antigens in the wbpB mutant). Full complementation of the wbpB mutation was observed only when multiple copies of flaA1 were provided using a complementation vector. This suggests that the reductase activity of FlaA1 is not very efficient, so that there is no actual functional redundancy in vivo. In addition, 4-keto sugar derivatives can be regarded as "pluripotent," since they can undergo various enzymatic modifications (52, 62, 71). Thus, the 4-keto-6-deoxy-UDP-GlcNAc generated by FlaA1 might be involved in multiple biosynthetic pathways and its reduction by WbpB might serve as a channeling mechanism to prevent its diversion toward other pathways. Because the wbpB gene is fairly rare in the bacterial world but is conserved and expressed in all H. pylori strains examined, the pathway at stake might be related to specific features of the biology and/or pathogenesis of H. pylori.
LPS structure (4, 5, 46) and biochemical data (15, 30, 36, 44) indicated that FlaA1 and WbpB were not LPS biosynthetic enzymes per se in H. pylori. Disruption of their genes nevertheless significantly reduced O-antigen production and resulted in the formation of altered core LPS (Fig. 4). In the absence of structural information for these mutants, it is reasonable to assume that band 1 observed on Tricine gels could correspond to the core and that the slower-migrating band 2 could correspond to the "core plus one O-chain unit." Band 3, which migrates only slightly slower than band 2, could represent a "core plus one" structure that carries an additional sugar residue on the terminal galactose of the O-chain unit. This additional sugar residue is likely to be L-fucose since the resulting difucosylated lactosamine motif is commonly found as a terminal structure in the H. pylori LPS (40, 46). This incorporation of an extra residue would explain the altered reactivity of the O chain produced by the wbpB mutant with the anti-Lewis Y antibody (Fig. 4). The activity of the transferase responsible for addition of this terminal sugar residue must be impaired or down-regulated in the flaA1 mutant so that no band 3 is present in this mutant and all O chains produced are of the Lewis Y type.
Both mutants were nonmotile (Fig. 7; Table 2), and the lack of motility correlated directly with the lack of flagellum production in the flaA1 mutant whereas the wbpB mutant still produced wild-type-like sheathed flagella. Western immunoblotting showed that flagellin(s) was produced by the flaA1 mutant, suggesting that it was the export or assembly of the flagellins into a flagellum that was impaired in this mutant. Flagellin O glycosylation has been found in other strains of H. pylori (34, 60) and has been proposed to be concomitant with its export to the bacterial surface (34). Considering our observation that the O-glycosylation sites described in the flagellin sequence of strain 1061 were conserved in the flagellins of strain NCTC 11637 and considering that glycosylation of surface appendage within one bacterial species is often strain dependent (3, 20, 59), we investigated if flagellins were also glycosylated in strain NCTC 11637. Our results obtained using enzymatic deglycosylation treatment and Western immunoblotting indicated that the flagellins of wild-type strain NCTC 11637 were not glycosylated. Thus, a defect in the posttranslational modification of the flagellins cannot be responsible for the lack of production of flagella in the flaA1 mutant or for the lack of functionality of the wbpB mutant flagella.
It has been reported previously that H. pylori mutants lacking the major flagellin, FlaA, do not produce flagella whereas mutants lacking the minor flagellin, FlaB, produce nonfunctional flagella (63). The possibility that FlaA would be missing in the flaA1 mutant and would prevent flagellum assembly cannot be excluded since the gel system and antibody used for these studies did not allow for discrimination between both flagellins. Similarly, the possibility that FlaB would be missing from the wbpB mutant could explain the presence of nonfunctional flagella. Alternatively, the lack of motility of the wbpB mutant might be due to a defect within a structural component of the basal body or motor of the flagella that has yet to be identified.
The defects in LPS production, membrane barrier properties, and flagellum-mediated motility associated with inactivation of flaA1 or wbpB suggested that both genes might be important for the virulence of H. pylori. In addition, the down-regulation of urease production observed in the flaA1 mutant (Fig. 2B and 9, bands a and e) might further compromise its survival in vivo, although the residual levels of urease produced were sufficient to sustain acid resistance in vitro (Fig. 3). Hence, FlaA1 and WbpB could represent valuable targets for the development of novel therapeutic agents against H. pylori. Since inactivating these enzymes abolishes the production of several virulence factors, it is unlikely that resistance to FlaA1- or WbpB-specific inhibitors could arise easily, in contrast to what is observed with currently available antibiotics (18, 31, 41). Mouse colonization assays are under way to investigate whether inactivation of flaA1 or wbpB results in defects in host colonization or survival in vivo.
The effect of a single gene on the synthesis of several virulence factors has been demonstrated previously (10, 20, 39) and involves direct incorporation of specific sugars within the structure of different virulence factors. In contrast, the sugars produced by FlaA1 (and WbpB) are not integral part of the virulence factors affected by disruption of either gene. We propose that these sugars are instead part of glycosylation motifs that target proteins involved in LPS or flagellum assembly and function and regulate their activity. Such regulation of protein function by their glycosylation has been observed previously in other bacterial species (17, 53, 55).
Several of the C. jejuni N-glycosylated proteins carry bacillosamine, a 2,4-diacetamido-2,4,6-trideoxyglucopyranose (72). Based on the biochemical function of FlaA1 (15) and considering the close relationship between H. pylori and C. jejuni, we propose that bacillosamine-dependent protein glycosylation also occurs in H. pylori and that FlaA1 is the first enzyme responsible for bacillosamine biosynthesis. While this work was in progress, it was shown that FlaA1 could complement the function of CJ1293, a homologue found in C. jejuni, and that CJ1293 was involved in the production of flagella (25). We also demonstrated that CJ1293 has the same biochemical activity as FlaA1 and performs a UDP-GlcNAc C-6 dehydration consistent with the first step of bacillosamine biosynthesis (12). Hence, these new data corroborate our hypothesis that FlaA1, like CJ1293, is involved in bacillosamine biosynthesis. Potentially glycosylated enzymes involved in LPS and flagellum synthesis are currently being analyzed in the context of the flaA1 mutation to confirm this hypothesis.
In summary, we demonstrate herein that FlaA1 and WbpB are functionally linked in the dual control of LPS biosynthesis and flagellum production and/or function in H. pylori and propose that this dual control involves protein glycosylation. This work is the first report to establish the existence of a regulatory network connecting the LPS and flagellum biosynthesis machineries and suggests that FlaA1- and WbpB-dependent protein glycosylation might be the underlying mechanism.
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ACKNOWLEDGMENTS |
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We thank D. E. Taylor (University of Alberta) and S. Logan (NRC, Ottawa, Canada) for providing strains 26695, NCTC 11637, and SS1 and R. Haas (Max von Pettenkofer Institute for Hygiene and Medicinal Microbiology, Munich, Germany) for providing plasmids pHel2 and pHel3. We thank J. S. Lam (University of Guelph) for support at the onset of this project and for providing anti-lipid A antibody, and we thank M. Smith and V. Somalinga for technical assistance. We thank D. Moyles (University of Guelph) for the electron microscopy analyses and A. Doherty-Kerby (University of Western Ontario) for MS analyses. Also, we also thank M. Valvano (University of Western Ontario) for the use of his Li-Cor Scanning system and S. Koval (University of Western Ontario) for the use of her microscope.
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FOOTNOTES |
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REFERENCES |
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, wild type plus pHel3; 
, flaA1 mutant; 
, wbpB mutant. (B) •, flaA1 mutant plus pHel2; 
, flaA1 mutant plus pHel2-wbpB. (C) 
, wbpB mutant plus pHel2; 
, wbpB mutant plus pHel2-wbpB; 
, wbpB mutant plus pHel2-flaA1.
