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1 Centre for Infection and Immunity, School of Medicine, Dentistry and Biomedical Sciences, Queen's University Belfast, Medical Biology Centre, 97 Lisburn Rd, Belfast BT9 7BL, UK
2 Institute of Cell Biology, University of Edinburgh, Darwin Building, Kings Buildings, Mayfield Road, Edinburgh EH9 3JR, UK
Correspondence
Garry W. Blakely
Garry.Blakely{at}ed.ac.uk
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). Capsule production in Escherichia coli is the paradigm for understanding the genetics and biochemistry of extracellular polysaccharide (EPS) production. In both E. coli and Salmonella enterica, polysaccharide biosynthetic genes are clustered together for the production of either the O-antigen of lipopolysaccharide (LPS) or capsular polysaccharides (groups 1, 2, 3 and 4; reviewed by Whitfield, 2006). Polysaccharides required for the production of LPS and capsules of groups 1 and 4 are transported across the inner membrane and polymerized by specific Wzx (flippase) and Wzy (polymerase) proteins, respectively. However, the fate of the polysaccharide is dictated by subsequent protein partners, which either ligate the polysaccharide to lipid A-core (WaaL for LPS and KLPS production) or transport it directly beyond the outer membrane (Wza for capsular polysaccharides). In contrast, group 2 and 3 capsular polysaccharides are synthesized by processive glycosyltransferases and exported via an ABC transporter system. Despite the large number of serotypes found in E. coli, there is generally little intra-strain variation in capsular types; for example, a strain will only produce a single group 1 capsule, and not multiple variants, because there is only one polysaccharide biosynthetic locus present in the chromosome. The exception is colanic acid, which may be co-expressed with group 2 capsules (Whitfield, 2006). Where intra-strain antigenic variation does occur, two different polysaccharides may be generated by modification of the polysaccharide sugar moieties, for example by acetylation (Vimr & Steenbergen, 2006).
In contrast, the genome of Bacteroides fragilis NCTC 9343 contains ten annotated regions (PS A–J) implicated, by homology, in EPS production (Cerdeño-Tárraga et al., 2005). PS loci A–I contain genes predicted to encode Wzx and Wzy proteins, suggesting that these polysaccharides will either be linked to lipid A-core prior to export or be secreted directly beyond the outer membrane. However, there are no annotated homologues for WaaL and only one potential Wza homologue can be found in the genome.
B. fragilis is the Gram-negative obligately anaerobic member of the normal human intestinal microbiota most frequently isolated from opportunistic infections (Patrick, 2002; Patrick & Duerden, 2006). Infections include peritonitis, serious gynaecological sepsis, soft tissue abscess and bacteraemia. The latter has an estimated mortality of 19 % (Redondo et al., 1995). Potential virulence determinants include expression of within-strain phase and antigenically variable polysaccharides that form a marginal electron-dense layer, or micro-capsule (MC), of approximately 35 nm in size, outwith the outer membrane and not visible by light microscopy (Fig. 1; Patrick et al., 1986; Lutton et al., 1991). Of the eight PS loci (A–H) that are related to variable MC expression, seven are switched ON and OFF by the site-specific inversion of promoter sequences, where recombination can be mediated by two members of the serine family of invertases (Patrick et al., 2003; Coyne et al., 2003; Liu et al., 2008). In addition to MCs, antigenically distinct and within-strain variable large capsules (LCs) and small capsules (SCs) are visible by light microscopy when cultures are grown in a glucose-defined medium (Fig. 1; Patrick et al., 1986), for which the gene loci remain to be assigned. Populations of B. fragilis enriched for the different capsules can be obtained by using Percoll density-gradient centrifugation. The antigenically variable MCs are co-expressed with the LC; an electron-dense layer adjacent to the outer membrane is visible beneath the LC by electron microscopy, and reactivity with MC-specific mAbs is demonstrable by immunofluorescence microscopy (Fig. 1; Lutton et al., 1991; Patrick, 1993). With continuous daily subculture these populations gradually revert to mixed capsular types. The LC confers resistance to phagocytic uptake and killing by human polymorphonuclear leukocytes in vitro (Reid & Patrick 1984), whereas MC-enriched populations are phagocytosed and killed. The MC is also important for colonization of the mammalian gastrointestinal tract. Recent evidence suggests that a single polysaccharide is sufficient to allow effective competition of a mutant strain with wild-type B. fragilis when co-inoculated into a gnotobiotic mouse (Coyne et al., 2008). This finding, however, is contradicted by other evidence that suggests strains expressing a single polysaccharide are not competitive in the gastrointestinal tract (Liu et al., 2008).
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). B. fragilis NCTC 9343 contains a shufflon that can generate eight different HsdS proteins that provide DNA target specificity for a type I R/M system, plus two further type I and two type III R/M systems (Cerdeño-Tárraga et al., 2005). Production of mutations in some strains of B. fragilis, often the 638R background, has been reported. One method relies on integration of the altered sequence into the chromosome followed by screening for spontaneous resolution of the diploid to yield the desired product (Coyne et al., 2003, 2008). This approach can be time consuming and does not always produce the required mutant if the diploid is stable. An alternative approach uses resistance to trimethoprim, conferred by a chromosomal thyA mutation in B. fragilis, to select for resolution of diploids where the suicide plasmid carries a functional copy of thyA (Baughn & Malamy, 2002). While this method enriches resolved diploids, it has the drawback that the strain to be engineered must first be made trimethoprim resistant and so all resulting strains contain a thyA mutation, which may have undesirable consequences for further studies. For example, a thyA mutation in E. coli will induce the SOS response if the cells are not provided with exogenous thymine (Begg & Donachie, 1978). Here we describe and validate a new and robust method for generating deletion mutants, and the complementation thereof, in B. fragilis NCTC 9343. This technique enables the rapid and reliable production of mutants that are either marked with an antibiotic resistance determinant or are markerless. The ability to complement non-polar deletions greatly facilitates genetic studies of this important human commensal and opportunistic pathogen. We also present direct evidence that two additional loci, PSJ and a region upstream of PSI, are involved in MC and LC biosynthesis respectively. These findings have implications for understanding how polysaccharides are synthesized in B. fragilis. |
METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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(Invitrogen) was used for constructing plasmids using standard molecular biology protocols. Conjugative vectors for introducing DNA into B. fragilis were based on pEP185.2 and were mobilized from the E. coli S17-1
pir strain (Simon et al., 1983). Plasmid pLyl01 (Li et al., 1995) was used to generate the I-SceI-expressing plasmid pGB920. Plasmid pVA2198 (Fletcher et al., 1995) was used to produce plasmids containing the locked ON and OFF promoter BF2782 constructs. Media for cultivation of B. fragilis included supplemented brain heart infusion (BHI-S) and defined medium (DM; van Tassell & Wilkins, 1978). Cultures were grown in a MACS MG-1000 or Min-MACS anaerobic workstation (Don Whitley), at 37 °C in an atmosphere of 80 % nitrogen, 10 % carbon dioxide and 10 % hydrogen. E. coli was cultured on LB at 37 °C.
Generation of deletion mutants.
Approximately 500 bp of DNA flanking the sequence to be deleted was amplified by PCR using Pfu polymerase (NEB). Flanking regions were fused to an ermF cassette by cross-over PCR using Pfu polymerase. PCR products were ligated into the multiple cloning site of pGB909 (a pEP185.2 derivative containing an I-SceI recognition sequence inserted at the SacI site). Ligated products were recovered in an E. coli S17-1pir strain. Constructs were mobilized from E. coli S17-1pir into B. fragilis NCTC 9343 using a filter-mating method (Valentine et al., 1988), followed by selection on BHI-S plates containing 10 µg erythromycin ml–1. Strains with integrated plasmids were confirmed by PCR and then electroporated with a derivative of pLyl01 containing the I-SceI coding region under the control of the B. fragilis fucR promoter, with selection for resistance to tetracycline. Transformants were streaked onto DM containing fucose and tetracycline. Colonies were then screened for resistance to erythromycin and deletion of the appropriate sequence confirmed by PCR. Markerless deletions were constructed by ligating cross-over PCR products from just the flanking regions into pGB910 (a derivative of pGB909 with an ermF cassette in the KpnI site). Conjugation and selection were as described above, except the final deletions were screened for loss of resistance to erythromycin.
Percoll density centrifugation.
Discontinuous Percoll gradients were prepared as previously described (Patrick & Reid, 1983). A 100 % stock Percoll (Pharmacia Biosystems) suspension was prepared by diluting sterile Percoll 9 : 1 with sterile 1.5 M sodium chloride, and adjusting the pH to 7.0 using 1 M hydrochloric acid. Percoll suspensions of 20 % and 40 % (v/v) were prepared by diluting the 100 % stock with sterile 0.15 M sodium chloride. Cells grown in DM were layered on top of the gradients followed by centrifugation at 2375 g for 40 min at 4 °C. Subculture of bacteria separating at and above the 0–20 % interface results in enrichment of the large capsulate population. Bacteria separating at the 20–40 % interface results in enrichment of the small capsulate population, whereas bacteria that pass further down the gradient are enriched for the MC.
Immunoblotting of EPS.
Immunoblots were prepared as previously described (Patrick et al., 2003). Aliquots of DM cultures, in late exponential phase, were resuspended in 1/4 diluted sample buffer [40 % (w/v) glycerol, 0.25 M Tris/HCl pH 6.8, 20 % (w/v) β-mercaptoethanol, 16 % (w/v) SDS and 0.04 % (w/v) bromophenol blue] and heated at 100 °C for 10 min. Then 25 µg proteinase K (Sigma) was added and incubated at 60 °C for 1 h. Digests were electrophoresed through Novex 10 % Tris/glycine polyacrylamide gels (Invitrogen) in Novex 1x Tris/glycine SDS running buffer (25 mM Tris base, 192 mM glycine and 0.1 % SDS pH 8.3). Polysaccharides were transferred to nitrocellulose membranes by electroblotting in transfer buffer [12 mM Tris base, 96 mM glycine, 20 % (v/v) methanol, pH 8.3]. Membranes were blocked for 1 h at 37 °C in blocking buffer [1x TBS, 5 % (w/v) dried semi-skimmed milk, 0.05 % (v/v) Tween-20], followed by washing in TBS-0.05 % Tween-20 (TBST) at 37 °C. Membranes were incubated with mAbs QUBf4 (Reid et al., 1987), 5, 6, 7, 8 (Lutton et al., 1991), 11 (Patrick et al., 1995), 18, 19 (Patrick, 1997), 25 (unpublished) and CE3 (Pantosti et al., 1995), for 1 h at 37 °C, followed by washing in TBST at 37 °C. The membrane strips were incubated for 1 h at 37 °C with either goat anti-mouse IgG (H+L) alkaline phosphatase conjugate (Bio-Rad) or goat anti-mouse IgM (µ chain specific) alkaline phosphatase conjugate (Sigma). The membranes were washed in TBST and polysaccharides visualized by detecting alkaline phosphatase activity using Sigma Fast BCIP/NBT (5-bromo-4-chloro-3-indolyl phosphate/nitro blue tetrazolium).
Transformation of B. fragilis NCTC 9343.
Overnight cultures of B. fragilis were diluted in pre-reduced BHI-S broth and grown, anaerobically at 37 °C, to an OD600 of 0.3. Cells were harvested by centrifugation at 1610 g for 20 min and washed five times in 20 ml pre-reduced ultrapure ice-cold water. Cells were finally resuspended in 1 ml pre-reduced ultrapure ice-cold water. One hundred microlitres of cells was used per electroporation in 0.2 cm electrode gene pulser cuvettes (Bio-Rad). If appropriate, diluted purified bacteriophage T7 Ocr (Overcomes classical restriction) protein, kindly gifted by Dr D. Dryden, School of Chemistry, University of Edinburgh, was added to the cells at the optimized concentration as detailed below. Then 500 ng plasmid DNA was added to the cells to give a final plasmid DNA concentration of 4.0 µg ml–1. Electroporation was performed using a Gene Pulser II electroporator (Bio-Rad) at 2.5 kV for 5–10 ms. Cells were resuspended in 1 ml pre-reduced 37 °C BHI-S broth and incubated anaerobically for 2–3 h at 37 °C prior to plating on BHI-S plates with appropriate antibiotics.
Capsule visualization.
B. fragilis possessing LC, SC and MC were identified using a negative capsule stain (Cruickshank, 1965). A 10 µl drop of culture was pipetted onto a glass slide. Five microlitres of carbol fuchsin (Sigma) was added to the bacterial suspension, mixed, and incubated at room temperature for 30 s. Then 5 µl eosin (Sigma) was applied and incubated at room temperature for 60 s. The stained cells were then smeared evenly across the slide using the edge of a microscope slide and air-dried at room temperature before visualization by microscopy. Cells and the background stained red, whereas capsules remain unstained. Transmission electron microscopy of ultrathin sections was carried out as previously described (Patrick et al., 1986).
Immunofluorescence microscopy (IFM).
IFM was performed as previously described (Patrick et al., 2003). Cultures of B. fragilis grown in DM broth or BHI-S broth were diluted in sterile PBS to an OD600 of 0.3. Then 10–30 µl was pipetted onto the wells of a 12-well Teflon-coated slide (ICN Biomedicals) and dried at 37 °C, followed by fixing with 100 % methanol at –20 °C for 20 min. Slides were brought to room temperature; mAbs were applied to each well and then incubated at 37 °C for 1 h. Slides were rinsed in a stream of sterile PBS and washed in a PBS bath for 20 min. Slides were removed and secondary antibody (goat anti-mouse IgG/IgM-FITC conjugate). Evans blue (0.1 %, Sigma), which stains proteins, was also added to each well as a counter-stain. Slides were then incubated for 1 h at 37 °C. The PBS washing step was repeated, and the slides were mounted in a glycerol-PBS antibleaching mounting fluid (Citifluor, Agar Scientific). Cells were examined using a Leitz Ortholux fluorescence microscope. Images were captured and analysed using an attached Nikon DMX 1200 digital camera and Lucia G/F software.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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to allow construction of deletions that are either marked by an antibiotic resistance determinant or are markerless. Marked deletion constructs are prepared by fusing 500 bp sequences from each side of the target gene, using PCR, to an intervening ermF cassette. The amplicon is ligated with an RP4-based conjugative vector (pEP185.2), into which we have inserted an ISce-I recognition sequence, prior to transformation of an appropriate E. coli host. The resulting strains are then conjugated with B. fragilis followed by growth in the presence of erythromycin to select for integration of the plasmid (Fig. 2). To resolve the diploid, the strains are transformed with a plasmid that expresses the ISce-I meganuclease under the control of the B. fragilis fucR promoter, followed by growth on a defined medium that contains fucose. Resolution of the diploid by homologous recombination follows the ISce-I-mediated double-strand break and results in production of either the wild-type or the deletion genotype (Fig. 2). Colonies are then screened for resistance to erythromycin and the presence of the deletion confirmed by PCR. The advantage of this method is that it enhances resolution of the diploid and so most transformants are either mutant or wild-type; this removes the need to repeatedly subculture diploid strains and screen large numbers of colonies for resolution by random recombination events.
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Transformation has not been previously reported for B. fragilis NCTC 9343, probably as a result of the multiple R/M systems specified by the genome. We therefore determined if the inclusion of purified T7 anti-restriction protein Ocr acted as a transformation-enhancing co-factor by inhibiting the action of the extensive endogenous R/M systems of B. fragilis. The effect of various concentrations of Ocr protein on the number of transformants was compared to the transformation efficiency in the absence of Ocr. Standard electroporations using 500 ng pVA2198 plasmid DNA per transformation, optimized in the absence of Ocr, generated 1.9x104 (±1.9x103) transformants per µg unmodified DNA. The number of transformants obtained following electroporation increased steadily with increased Ocr protein concentration, reaching between 5.0x105 (±2.5x104) and 5.2x105 (±2.2x104) transformants µg–1, corresponding to 16 µg and 8 µg Ocr protein per plasmid transformation. When Ocr concentration was increased from 16 µg to 32 µg protein per transformation, a tenfold decrease in transformant numbers was observed. An extensive selection of negative controls and transformant identification tests were performed, confirming that all erythromycin-resistant colonies identified were B. fragilis NCTC 9343 transformed with plasmid pVA2198. Therefore 16 µg purified T7 Ocr anti-restriction protein (127 µg ml–1) was used per transformation. It has been shown that Ocr protein is capable of inhibiting type I R/M systems covering a diverse range of eubacteria and archaea, indicating that the anti-restriction property of the protein is not dependent on the target DNA sequence of the type I R/M enzymes (Krüger et al., 1977, 1983; Walkinshaw et al., 2002), as each type I system recognizes a unique target DNA sequence (Titheradge et al., 2001). It is likely that the negative charge of this DNA mimic allows electroporation of the protein into the cell, where it is able to reduce the activity of the resident type I R/M systems (Hoffman et al., 2002; Dryden, 2006). The all-encompassing type I anti-restriction activity of Ocr may therefore explain why the protein appears to be capable of improving transformation efficiency in B. fragilis NCTC 9343 by counteracting the DNA-degrading effects of all three complete type I R/M systems, including the phase-variable system.
Validation of the deletion technique by disruption of polysaccharide gene cluster PSC
To validate our deletion strategy, we chose to replace the first gene in PSC, upcY, with an ermF cassette. Krinos et al. (2001) had previously shown that disruption of upcY, by insertion of a non-replicating plasmid, prevented expression of the associated polysaccharide. Expression of PSC is not controlled by an invertible promoter and its regulatory mechanisms are currently unknown. However, the protein encoded by upcY is predicted to contain a KOW domain and has homology to NusG-like proteins; it is therefore likely to be involved in anti-termination of transcription within the leader sequence of the polysaccharide cluster.
We analysed the 
upcY : : ermF strain by immunoblotting, IFM and negative staining of capsules using a collection of mAbs that specifically bind to distinct polysaccharides produced by B. fragilis NCTC 9343 (Patrick et al., 2003; Fig. 3). Polysaccharides from the wild-type and mutant strains were prepared by proteinase K digestion of whole cells followed by electrophoresis through 12–20 % polyacrylamide gels. Immunoblots, using all mAbs, allowed detection of characteristic heteropolymorphic, high-molecular-mass polysaccharides (HMMPS) produced by the parental strain, as described previously (Lutton et al., 1991). Polysaccharides derived from the upcY : : ermF strain were detectable on immunoblots using eight of the mAbs, but not QUBf7 (Fig. 3a). Binding of this mAb has previously been associated with polysaccharides produced by gene cluster PSC (Coyne et al., 2000). IFM using the same mAbs confirmed that deletion of upcY led to an inability to detect cell-surface-associated polysaccharides that reacted with QUBf7; however, expression of other polysaccharides was unaffected (Fig. 3b). Both the LC and SC could be detected by negative staining of capsules (data not shown), indicating that only production of the PSC MC was affected by the mutation. It is possible that insertion of ermF has a polar effect on transcription, which subsequently leads to this phenotype. Together these data confirm that deletion of upcY, and its replacement with ermF, abolishes expression of the PSC gene cluster and illustrate the effectiveness of the allelic replacement methodology.
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; Lindberg et al., 1990, Lutton et al., 1991). Dispersed in the NCTC 9343 genome are a number of ‘orphan’ genes that encode putative polypeptides with homology to proteins known to function in EPS biosynthesis. The original annotation of the genome identified two potential members of the polysaccharide copolymerase (PCP) family, encoded by BF1708 and BF2784 (Cerdeño-Tárraga et al., 2005). No other proteins encoded by B. fragilis NCTC 9343 can be identified as PCPs using BLAST searches (E-value cutoff 3x10–4). The assignment of these two proteins as PCPs is supported by their current inclusion as the only proteins in B. fragilis within the InterPro003856 family, which contains 1897 members. PCPs are divided into three classes based on function and common secondary structural features (Morona et al., 2000). Gene BF2784 encodes a polypeptide with homology to proteins of the PCP2 class, for example Wzc proteins that are involved in capsular biosynthesis (Whitfield, 2006). Gene BF1708, within PS gene cluster J/10, encodes a putative member of the PCP1 class, which includes Wzz proteins involved in O-antigen synthesis. Members of this family often show limited homology but have predicted N- and C-terminal transmembrane helices separated by a periplasmic coiled-coil region (Tocilj et al., 2008). The polypeptide encoded by BF1708 has 20 % identity to E. coli WzzB and is predicted to contain the secondary structure features associated with PCP1 members. In E. coli the Wzz protein modulates the polymerase activity of the Wzy involved in LPS biosynthesis to produce a distinctive modal distribution of O-antigen chain lengths (Franco et al., 1998). Although BF1708 encodes the only identifiable homologue of Wzz in the NCTC 9343 genome, the high level of sequence divergence within the PCP classes means that other members might also be present. To address the nature of the polysaccharide associated with LPS in B. fragilis and determine if the protein encoded by BF1708 was involved in LPS biosynthesis, we deleted the gene by replacing it with an ermF cassette and then analysed the polysaccharides produced by the mutant. The introduction of the ermF cassette could have a polar effect on expression of two downstream genes, BF1709 and BF1710; however, the products of these two genes are not likely to be central to polysaccharide production and are annotated as a putative transcriptional regulator and a protein of unknown function but with similarity to a dTDP-4-dehydrorhamnose 3,5-epimerase.
The first obvious phenotype of the 1708 strain was the bridging flocculation of cells during culture in liquid medium (Fig. 4a), which may be indicative of altered cell-surface properties. Microscopy with negative staining for capsules revealed bacteria with either LC or SC, either individually or associated with the aggregates composed of bacteria non-capsulated by light microscopy. Labelling with a mAb specific for the SC confirmed its presence both by fluorescence microscopy and immunoblotting of polysaccharides (not illustrated). Examination of the capsule smears revealed an irregular capsular phenotype in approximately 10 % of the LC cells (Fig. 4b). This phenotype has not been observed previously in B. fragilis and how it relates to the deletion of BF1708 is open to speculation. Immunoblotting of MC polysaccharides produced by the 1708 strain indicated that this culture was not antigenically mixed but was predominantly expressing a single polysaccharide that reacted with mAb QUBf5, which is specific for polysaccharides synthesized by the PSD locus (Fig. 4c). The blot pattern for the polysaccharide was, however, clearly distinct from that of the parental strain and other mutants, such as the upcY strain, which show HMMPS and an associated ladder similar to that previously described (Lutton et al., 1991; Fig. 4c); in the 1708 mutant the HMMPS was absent and a lower-molecular-mass ladder pattern was evident (Fig. 4c). The loss of HMMPS D in the 1708 strain is supported by the reduction in reactivity with mAb QUBf5 by IFM (Fig. 4d). The inability to detect HMMPS produced by the other loci, using immunoblotting and IFM, does not rule out the possibility that the polysaccharides are still present as short-chain repeats, which would reflect a reduced sensitivity of the mAbs for the residual repeat units. Together, these data indicate that deletion of the putative wzz gene and replacement with ermF prevents assembly of HMMPS synthesized by the PSD gene cluster into a long-chain-length polymer attached to the cell surface and that other HMMPS associated with the MC are not detectable.
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when they described the separation of bacteria expressing capsules of different sizes using centrifugation through Percoll step gradients. While expression of seven EPS associated with the MC are regulated by site-specific recombination, little is known about the genes or the regulation of LC or SC production; clearly from the data presented above, the putative Wzz encoded by BF1708 is not involved. Annotation of the NCTC 9343 genome, however, identified a further set of orphan genes which are potentially involved in polysaccharide biosynthesis. ORFs BF2782 and BF2784 may encode homologues of the WbaP and Wzc proteins, which are involved in group 1 and 4 capsule synthesis in E. coli. WbaP is part of a glycosyltransferase complex involved in formation of lipid-linked precursors in the cytoplasm prior to translocation across the inner membrane, while Wzc (a member of the PCP2 family) is a membrane-bound protein that modulates the activity of the cognate Wzy polymerase. The promoter upstream of BF2782 was reported to be invertible and its orientation was reported to correlate with production of the LC, as shown by deletion of a tyrosine site-specific recombinase, tsr19 (Chatzidaki-Livanis et al., 2008). These authors, however, cultured their isolates in a complex basal medium that fails to reveal the LC phenotype and therefore does not distinguish between the antigenically different LC and SC. The LC and SC phenotypes are clearly evident on culture in defined medium (DM) by light microscopy (Fig. 1
; Patrick & Reid, 1983; Patrick et al., 1986) and are antigenically distinct (Reid et al., 1987). LC and SC bacteria can be separated using a discontinuous Percoll density gradient where the LC population does not enter into the 20 % Percoll, but remains at the top of the gradient after centrifugation, whereas the SC cells locate at the 20–40 % interface layer (Fig. 5a) The MC cells pass through the gradient to the 60–80 % interface (Patrick et al., 1986). Capsule staining of wild-type cells from the 0–20 % interface shows enrichment of cells expressing the LC, which are visually distinct from the SC population derived from the 20–40 % interface (Fig. 5a). The Percoll gradients used to fractionate the tyrosine site-specific recombinase mutant illustrated by Chatzidaki-Livanis et al. (2008) show that the mutant with the ‘locked ON’ promoter has entered into the 20 % Percoll suspension, and therefore it is not clear whether the LC or SC has been affected by the deletion. The negative staining of cells with the locked ON promoter reported by Chatzidaki-Livanis et al. (2008) also did not display a capsule that correlated with the previously described characteristics of the LC (Patrick & Reid, 1983; Patrick et al., 1986; Reid et al., 1987; Lutton et al., 1991). To determine directly the role of BF2782 in production of either the LC or SC, we therefore generated both marked and markerless deletions of the gene. Replacement of BF2782 with an ermF cassette resulted in a mutant that was unable to synthesize the LC, as shown by the failure of cells to remain on top of 20 % Percoll after centrifugation. This was confirmed by capsule staining of bacteria grown in DM followed by microscopy, which failed to detect any cells producing the LC (data not shown). IFM (Fig. 5b) and PAGE followed by immunoblotting (data not shown) of the BF2782 mutant using a suite of MC-specific mAbs indicated that deletion of BF2782 had not affected expression of the multiple-variable MC. Since insertion of ermF may have been polar, we analysed the strain containing a markerless deletion of BF2782 using the same methods. The markerless mutant did not produce cells that remained on top of the 20 % Percoll after centrifugation (Fig. 5c), and again we could not detect any cells expressing the LC by capsule staining (data not shown). SC bacteria were, however, evident at the 20–40 % interface (Fig. 5c), and reaction with a mAb specific for the SC (Reid et al., 1987) was positive by IFM (Fig. 5b). To confirm that the markerless deletion was responsible for the loss of LC production, we complemented the mutation by expressing BF2782 from a plasmid-borne copy of the gene. Two plasmids containing the BF2782 ORF were constructed: one had the invertible promoter locked ON while the other was always OFF. Only transformation of the 2782 strain with the ON promoter construct enabled the cells to produce the LC, as determined by their presence on the 0–20 % Percoll interface (data not shown) and capsule staining (Fig. 5d). Expression of the LC in this strain had also returned to being phase variable, as only a fraction of the population produced the LC. This observation indicates that the invertible promoter on the chromosome of the markerless 2782 strain was switching the expression of the remaining downstream gene(s) on and off, and implies that either BF2783 or BF2784, or both, are also involved in LC production.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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). It is clearly different from the paradigm of classical enteric LPS; for example, lipid A diglucosamine in B. fragilis is monophosphorylated rather than bisphosphorylated. While all the genes necessary to synthesize keto-deoxyoctonate (KDO) are present in the genome of NCTC 9343, its chemical composition is different from enteric KDO since it appears to be phosphorylated, thus rendering it undetectable in the standard thiobarbituric acid assay (Beckmann et al., 1989). A major question, however, remains over the presence or absence of a polysaccharide that resembles an enteric O-antigen. Lindberg et al. (1990) stated that the repeating unit of an O-antigen is not present in B. fragilis, while Poxton & Brown (1986) clearly demonstrated that a polysaccharide with a repeat size equivalent to enteric O-antigen could be detected in at least one B. fragilis strain. Subsequently Lutton et al. (1991) described a within-strain antigenically variable HMMPS with an associated O-antigen-like ladder pattern after PAGE and immunoblotting, which was later designated PSD (Patrick et al., 2003).
The genome of B. fragilis NCTC 9343 appears to be devoid of an operon with genetic structure similar to the loci involved in synthesis of O-antigen or lower-molecular-mass lipid A-linked K-antigen (termed KLPS) in E. coli. However, a single gene encoding a protein with homology to an O-antigen copolymerase, Wzz, was identified in B. fragilis. The Wzz protein in E. coli determines the modal distribution of O-antigen/KLPS chain length, and deletion of the wzz gene simply produces unregulated lengths of polysaccharide chains. In group 1 and 4 K-antigens, for example, serotype K40, which forms both lipid A-linked KLPS and high-molecular-mass non-lipid A-linked capsular (K)-antigen, there is no loss of higher-molecular-mass K-antigen when wzz is deleted. Deletion of BF1708 produced an unexpected phenotype whereby the HMMPS representing the MC were not detectable by immunoblotting using mAbs. This phenotype is similar to that described by Coyne et al. (2008) following deletion of a gene encoding a UDP-GlcNAc 4,6-dehydratase (BF1706), where the strain failed to produce polysaccharides associated with the MC and also underwent aggregation. It is hypothesized that the dehydratase is responsible for synthesizing the unusual sugar residues found in B. fragilis polysaccharides. Although both BF1706 and BF1708 seem to be involved in HMMPS biosynthesis, it is not clear if they are part of an operon since BF1706 is separated from BF1707 by 120 bp.
The loss of HMMPS in the B. fragilis wzz/1708 mutant is very different from the change in distribution of O-antigen chain lengths in an E. coli wzz mutant, which was shown to be random and contained both larger and smaller numbers of repeat units compared to the wild-type (Franco et al., 1998). The demonstration that a Wzz homologue is involved in controlling polysaccharide chain length also suggests that PSD and the other MC-associated polysaccharides may be covalently linked to lipid A-core. In group 1 or 4 capsules, lipid A-linked KLPS usually only represents a small proportion of total capsular polysaccharide, with most being anchored to an unknown moiety on the cell surface as capsular or K-antigen. This would therefore make the antigenically variable MC-associated polysaccharides the equivalent of enteric LPS, but with a higher-molecular-mass polysaccharide that forms the electron-dense layer visible by electron microscopy on the exterior of the cell. Some strains of E. coli do produce very long-chain-length O-antigen polysaccharides, for example E. coli O157 (Sheng et al., 2008); however, these polymers are still shorter than the HMMPS present in B. fragilis MC. A high-molecular-mass LPS in B. fragilis would be consistent with the findings of Lindberg et al. (1990). We also hypothesize that the one B. fragilis strain (NCTC 9344), which Poxton & Brown (1986) found to express LPS with short repeat units, potentially contained a mutation in wzz. This would be consistent with the short polymer repeat units that we observed on the immunoblots of the wzz/1708 strain. Whether or not the MC polysaccharide is anchored to the lipid A, and therefore equivalent to a high-molecular-mass LPS, could be tested by deleting the gene encoding the ligase that attaches the polysaccharide to lipid A-core; however, homologues of waaL have yet to be identified in the genome. Nevertheless, the MC polysaccharide is present in LPS prepared using, for example, aqueous phenol extraction methods (Delahooke et al., 1995).
Three different capsule types were originally identified in B. fragilis: MC, LC and SC (Patrick et al., 1986). If the MC is anchored to lipid A-core and equivalent to a high-molecular-mass KLPS, then are the LC and SC equivalent to E. coli groups 1 and 4, and related colanic acid, capsules? Association of mAb binding with expression of specific polysaccharide gene clusters has narrowed the focus to potential candidate genes involved in LC and SC formation. The genome sequencing project annotated three genes (BF2782–2784) encoding homologues of proteins potentially involved in group 1 and 4 capsule synthesis (Cerdeño-Tárraga et al., 2005). An invertible promoter has been identified upstream of these three genes, and deletion of the tyrosine recombinase that mediates inversion of the promoter has allowed expression to be locked either in the ON or OFF conformation (Chatzidaki-Livanis et al., 2008). The OFF position correlated with loss of capsule visible by light microscopy. These authors, however, used a complex medium in which the LC is not distinguishable from the SC by light microscopy or Percoll gradient centrifugation (Patrick & Reid, 1983). We tested directly the hypothesis that BF2782 was involved in LC formation by deleting the gene and then complementing the mutation with a plasmid-borne copy of the gene. Deletion of BF2782 prevented the synthesis of the LC as determined by Percoll gradient centrifugation and capsule staining. The SC, however, was still expressed and this was confirmed by mAb labelling. Complementation of the non-polar markerless deletion resulted in phase-variable expression of the LC, even when BF2782 was constitutively expressed. This observation suggests that inversion of the promoter controlling expression of BF2783 and BF2784 in the 2782 strain was responsible for switching OFF expression and therefore implicates these genes in expression of the LC. The predicted protein encoded by BF2784 has homology to Wzc, which modulates the polymerase activity of the Wzy protein, and BF2783 has homology to Wza, a member of the outer-membrane auxiliary (OMA) protein family, which forms multimeric channels involved in polysaccharide extrusion. From these data we infer that assembly of the LC in B. fragilis is equivalent enzymically to production of colanic acid, group 1 and 4 K-antigen capsules in E. coli, whereas the MC are assembled by an entirely independent mechanism similar to that of KLPS. B. fragilis is therefore unusual not just in the number and variety of different polysaccharide capsule loci present in an individual strain, but also potentially in the nature of the LPS anchored at the cell surface. While the role of the LC in avoidance of phagoctyic uptake and killing and the MC in resistance to serum killing has been reported (Reid & Patrick, 1984), the relative importance of these structures and the SC in survival in the human gastrointestinal tract remains to be determined.
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ACKNOWLEDGEMENTS |
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Edited by: V. Eijsink
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Received 23 October 2008;
revised 19 December 2008;
accepted 19 December 2008.
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) and resolution of the diploid by generation of either the deletion or wild-type genotypes by homologous recombination. Chromosomal DNA is indicated as dashed black lines; plasmid DNA is indicated as solid grey lines.
