Helicobacter pylori lipopolysaccharide modification, Lewis antigen expression, and gastric colonization are cholesterol-dependent
© Hildebrandt and McGee; licensee BioMed Central Ltd. 2009
Received: 16 June 2009
Accepted: 14 December 2009
Published: 14 December 2009
Helicobacter pylori specifically takes up cholesterol and incorporates it into the bacterial membrane, yet little is currently known about cholesterol's physiological roles. We compared phenotypes and in vivo colonization ability of H. pylori grown in a defined, serum-free growth medium, F12 with 1 mg/ml albumin containing 0 to 50 μg/ml cholesterol.
While doubling times were largely unaffected by cholesterol, other overt phenotypic changes were observed. H. pylori strain SS1 grown in defined medium with cholesterol successfully colonized the stomach of gerbils, whereas SS1 grown without cholesterol failed to colonize. H. pylori lipopolysaccharide often displays Lewis X and/or Y antigens. Expression of these antigens measured by whole-cell ELISA was markedly enhanced in response to growth of strain SS1, 26695, or G27 in cholesterol. In addition, electrophoretic analysis of lipopolysaccharide in wild type G27 and in mutants lacking the O-chain revealed structural changes within the oligosaccharide core/lipid A moieties. These responses in Lewis antigen levels and in lipopolysaccharide profiles to cholesterol availability were highly specific, because no changes took place when cholesterol was substituted by β-sitosterol or bile salts. Disruption of the genes encoding cholesterol α-glucosyltransferase or lipid A phosphoethanolamine transferase had no effect on Lewis expression, nor on lipopolysaccharide profiles, nor on the cholesterol responsiveness of these properties. Disruption of the lipid A 1-phosphatase gene eliminated the effect of cholesterol on lipopolysaccharide profiles but not its effect on Lewis expression.
Together these results suggest that cholesterol depletion leads to aberrant forms of LPS that are dependent upon dephosphorylation of lipid A at the 1-position. A tentative model for the observed effects of cholesterol is discussed in which sequential steps of lipopolysaccharide biogenesis and, independently, presentation of Lewis antigen at the cell surface, depend upon membrane composition. These new findings demonstrate that cholesterol availability permits H. pylori to modify its cell envelope in ways that can impact colonization of host tissue in vivo.
Helicobacter pylori is a highly niche-adapted pathogen that inhabits the human stomach, is transmitted primarily within families, and has no known environmental reservoir. Chronic infections may be asymptomatic or cause gastritis, ulcer, or gastric cancer. To establish infection, the bacterium must survive transit through the acidic gastric compartment . It penetrates and establishes residence in the protective mucus layer, a lipid- and cholesterol-rich environment [2, 3]. Within this niche the bacterium employs a variety of mechanisms to evade host immune response.
Lipopolysaccharides (LPS) on the surface of H. pylori are modified to display certain human blood group antigens, primarily Lewis antigens X and Y [4–7], and less frequently H type 1, i-antigen, blood group A, or Lewis antigens A or B [8–10]. These surface LPS antigens are necessary for the establishment of infection, because mutant strains defective for LPS O-antigen synthesis or for Lewis X/Y expression fail to colonize mice [11–13]. There is evidence that Lewis antigens expressed on the bacterial surface contribute to adherence of H. pylori to gastric epithelial cells [10, 14], and play a role in tissue tropism [15–17]. Gastric epithelial cells also express Lewis antigens [18, 19], suggesting that the display of Lewis antigens on the bacterial surface may serve as a mimicry strategy. Studies of clinical isolates [18, 20] and experimental infections in animals  support this role for bacterial Lewis antigens in immune evasion. In human infection, H. pylori Lewis antigens have been linked to the severity of peptic ulcer and duodenitis [16, 22]. Another important feature of H. pylori LPS is its modified lipid A structure, with reduced acylation and fewer charged groups than is typical of enterobacteria . These lipid A modifications minimize endotoxic and inflammatory properties of H. pylori LPS (reviewed in ).
Cholesterol is a nonessential nutrient for H pylori, though it promotes growth in serum-free media [25, 26]. H. pylori specifically incorporate cholesterol into the bacterial membrane , as do a limited number of pathogenic and commensal bacteria including Proteus mirabilis, Lactobacillus acidophilus, Borrelia sp., and Mycoplasma [28–30]. Cholesterol may strengthen the membrane in these organisms [30–32]. H. pylori also uniquely form cholesterol α-glycoside [33, 34], and this metabolite can be further modified by acylation or phosphatidylation . Alpha-glucosylated cholesterol subverts host immune response to the bacterium in a mouse model, through suppression of phagocytosis and of T cell activation . Other roles for cholesterol and cholesterol metabolites in the bacterial membrane have yet to be explored. In this report, we demonstrate that the biosynthesis of lipopolysaccharide, including Lewis antigen expression and LPS core/lipid A modification, are altered by availability of cholesterol in the growth medium. We present data indicating that these changes in the cell envelope may significantly influence the pathogen/host interaction in an animal model of infection.
Bacterial strains and growth conditions
Strains of H pylori included the laboratory strain ATCC43504 (origin: Australia), 26695 (UK), clinical isolate G27 (Italy , provided by N. Salama), and the mouse adapted strain SS1 (Australia; provided by Adrian Lee ). Bacteria were maintained at 37°C in a microaerobic atmosphere of 5% O2/10% CO2 on Campylobacter blood agar (CBA). Bacteria were passaged every 2 to 3 days, and for no more than 25 days, to minimize genetic drift. For growth in chemically defined medium , bacteria were inoculated from CBA into tissue culture flasks containing Ham's F12 (Gibco) with 1 mg/ml bovine serum albumin (fatty acid-free, Sigma A7906), referred to throughout as defined medium. Liquid cultures were passaged daily by dilution into fresh medium at initial densities of 1-2 × 106/ml, and used at passage 3 to 5. Cell culture grade cholesterol (>99%, Sigma) was added to F12 as a stable 10× emulsion containing 500 μg/ml cholesterol dispersed in 10 mg/ml albumin, which was prepared according to . The following media additions were carried out in like manner: β-sitosterol (synthetic, 95%), sodium taurocholate, sodium glycocholate, β-estradiol, progesterone (all from Sigma), dehydroepiandrosterone (Calbiochem), and β-coprostanol (Matreya).
Doubling times were determined during log phase growth by quantitating viable cells using the Cell Titer Glo reagent (Promega) as validated and described . Measurement of biomass as CFU, as cellular protein, or as ATP have all produced consistent results. A value of 1 attomol ATP per cell  was assumed for routine passage. Possible inaccuracy of this value does not fundamentally influence interpretation of data.
primers for allelic disruption a
CAT fwd 
CAT rev 
additional primers for confirmation of gene disruption
Animal experiments were approved by the LSUHSCS Institutional Animal Care and Use Committee. Female Mongolian gerbils were maintained on ordinary diet ad libitum. To preserve motility, H. pylori strain SS1 was cultured overnight under microaerobic conditions in T75 flasks containing 40 mls of F12 medium with 0.4 mg/ml albumin and 0 or 50 μg/ml cholesterol. The motile planktonic bacteria were harvested by centrifugation and resuspended in isotonic saline. Colony forming units (CFU) were measured in these inocula by serial dilution and plating on CBA, and these measurements confirmed equal dosage of viable bacteria between the two growth conditions. Approximately 108 CFU per 30 μl were given orally to animals (n = 6 to 9 per group). Animals were euthanized 11 days later, and stomachs were removed and dissected. H. pylori present in gastric antrum homogenates were quantitated by serial dilution and plating on CBA containing 5-fluorocytosine (5 μg/ml), vancomycin (10 μg/ml), amphotericin B (5 μg/ml), bacitracin (30 μg/ml), polymyxin B (10 U/ml), and trimethoprim (10 μg/ml) . Duplicate CFU determinations were made for multiple dilutions of each tissue sample.
Standard procedures [6, 7, 45], were adapted for the use of peroxidase conjugated secondary antibody. All antibodies were obtained from Calbiochem. Overnight cultures of bacteria were collected by centrifugation at 3500 × g for 10-15 min, washed in Dulbecco's phosphate buffered saline, and repelleted at 10,000 × g for 2 min, then resuspended in 15% glycerol/0.9% NaCl. The cell suspensions were assayed for protein content and stored at -20°C. Cell samples containing known amounts of protein were rapidly diluted into 50 mM sodium bicarbonate/carbonate pH 9.55 and dispensed immediately into wells of an ELISA plate (Costar #9017). Plates were sealed and refrigerated overnight, then blocked for 90 min in 3% bovine serum albumin dissolved in the wash buffer which consisted of 0.1 M sodium phosphate pH 7.4/0.1 M NaCl/0.1% w/v Tween-20. Primary antibody, monoclonal anti-Lewis X (Signet clone P12) or anti-Lewis Y (Signet clone F3), diluted 1:500 in wash buffer/1% BSA, was added for 2 hours, followed by four changes of wash buffer. The secondary antibody, a 1:2500 dilution of horseradish peroxidase-conjugated goat anti-mouse IgM in wash buffer/1% BSA, was added for 90 min, followed by four changes of wash buffer. The chromogenic substrate was 0.42 mM tetramethylbenzidine and 0.02% H2O2 in 50 mM acetate/citrate pH 5.5 . After 15 minutes at room temperature, reaction was stopped with 1/5th vol 2.5 N H2SO4, and color change was measured in a plate reader at 450 nm. In negative controls omitting either primary or secondary antibody, or with E. coli strain HB101 substituted for H. pylori, color change was negligible (A<0.05). Levels of Lewis Y were negligible (A<0.1) in strain 26695 or 43504, as were Lewis X levels in SS1.
Electrophoretic analyses of lipopolysaccharides
H. pylori cultures were collected as described above, and washed cell pellets were stored at -70°C. Cells were lysed in 60 mM Tris HCl pH 6.8 containing 2% SDS at 95-98°C for 10 min. Protein content was measured using the bicinchoninic acid assay (Pierce). Samples of cell lysates were adjusted to equal protein content (1 mg/ml), then proteolyzed in reactions containing (final) 60 mM Tris HCl pH 6.8, 0.67% SDS, and 0.67 mg/ml proteinase K at 60°C for 2 hours . To eliminate electrophoretic artifacts due to the presence of lipid/detergent complexes, proteolyzed samples were extracted with hot phenol . Control experiments verified that all LPS bands were recovered through the following extraction procedure qualitatively and without bias. Proteolyzed samples were mixed with 1 volume of 90% aqueous phenol and incubated at 70°C for 20 min. After cooling to 10°C for 1 min, the samples were centrifuged at 12,000 × g for 20 min at 10°C, and the aqueous phase collected. The phenolic phases were re-extracted with 1 volume of H2O at 70°C for 10 min, and the centrifugation repeated. The combined aqueous extracts were adjusted to 0.5 M NaCl and precipitated with 10 vol ethanol in the refrigerator overnight, then centrifuged at 20,000 × g for 20 min at 10°C and air dried. Purified LPS samples were redissolved in Laemmli sample buffer  at 95°C for 5 min. Samples were applied to 15% polyacrylamide/0.9% bis minigels containing 3.2 M urea with the Laemmli discontinuous buffer formulation , and a 5% stacking gel. After electrophoresis at 150 V for 75 min, gels were either fixed overnight for silver staining  or transferred to polyvinylidenedifluoride membrane using Tris/glycine transfer buffer . Blots were blocked overnight in 3% bovine serum albumin and 0.03% NaN3 in the wash buffer described above for ELISA. Primary antibody (anti-Lewis X or anti-Lewis Y, 1:200) and secondary antibody (peroxidase-conjugated goat anti-mouse IgM, 1:1000) were diluted in wash buffer containing 0.5% BSA. Colorimetric detection used 3,3'-diaminobenzidine with cobalt enhancement . Densitometry was performed with the public domain application Image J, available at http://rsb.info.nih.gov/ij.
Enhancement of cell surface Lewis antigen expression by the growth of cultures in the presence of cholesterol.a
fold increase compared to parallel cholesterol-free culture
mean ± SEM (n)
mean ± SEM (n)
4.32 ± 0.36 (6)
1.88 ± 0.08 (5)
G27 wild type
2.85 ± 0.42 (8)
2.22 ± 0.24 (8)
3.69 ± 0.34 (5)
2.88 ± 0.30 (5)
2.59 ± 0.50 (6)
2.47 ± 0.43 (7)
Enhanced cell surface Lewis antigen expression is cholesterol-specific
fold increase compared to parallel cholesterol-free culture
mean ± SEM (n)
mean ± SEM (n)
2.96 ± 0.22 (5)
2.48 ± 0.10 (4)
1.80 ± 0.47 (4)
1.19 ± 0.13 (3)
0.64 ± 0.16 (4)
0.84 ± 0.20 (3)
We also investigated cholesterol responsiveness of LPS in a G27 pmi::cat strain lacking O-antigen chains (Figure 9B). As in wild type G27, this strain showed the presence of an additional, more slowly-migrating band in the core region that was diminished or lost upon growth in cholesterol. Likewise, pmi::cat strains of 26695 and SS1 also lacked O-chains, and also exhibited similar cholesterol-dependent band loss in the conserved LPS core region (data not shown). Since LPS species migrating in this region likely include only core oligosaccharide and lipid A moieties, we directed our attention to these components in trying to identify specific cholesterol-dependent structural modifications. We selectively disrupted two lipid A modification genes, either lpxE or eptA, encoding the lipid A 1-phosphatase and lipid A phosphoethanolaminetransferase, respectively . Then, LPS profiles were compared in pairwise cultures of these mutated G27 strains grown in the presence or absence of cholesterol (Figure 9C). We found that the eptA::cat strain retained an LPS response to cholesterol that was even more distinct than in the wild type. In contrast, cholesterol-responsive bands were abolished in the lpxE::cat strain. These results implied that the aberrant bands which accumulated under conditions of cholesterol depletion in the wild type, but not in lpxE::cat, may represent forms of LPS in which the lipid A moiety has been dephosphorylated at the 1-position. It is also possible that, in these bands, the core may have undergone further modification subsequent to lipid A dephosphorylation (see Discussion).
In eukaryotic membranes, cholesterol modulates curvature and fluidity, and cholesterol-rich lipid subdomains influence numerous membrane functions, including signal transduction and transport activity , yet very little is known about the physiological roles of cholesterol among the prokaryotes that utilize it. In this study, we used chemically defined medium to begin to characterize these roles of cholesterol in H. pylori. Growth of H. pylori in the presence of cholesterol proved to be essential for gastric colonization in the gerbil, even though it is not necessary for growth in vitro. This colonization experiment was conducted under standard dietary conditions, where cholesterol should be abundant in gastric mucus [2, 3, 60]. Taking into account that H. pylori can also acquire cholesterol from the membrane of host gastric epithelial cells , our data would suggest that incorporation of cholesterol into the bacterial membrane prior to inoculation may facilitate early steps in gastric colonization that precede adherence to host epithelium, such as motility and/or acid resistance. Preliminary experiments have indicated that H. pylori grown in the presence of cholesterol are more resistant to acid and oxidative stresses than when cholesterol-depleted (DJM, unpublished observations). We propose that incorporation of cholesterol and/or cholesterol metabolites may strengthen the bacterial membrane against such stresses, protecting the bacterium from gastric acid prior to entry into the more pH-neutral gastric mucus layer. Once the epithelial layer has been colonized, host-derived cholesterol may then be utilized.
We have also presented evidence of a role for cholesterol in establishment of the normal lipopolysaccharide component of the cell envelope. Both Lewis antigen[12, 14] and core oligosaccharide [13, 61, 62] contribute to H. pylori adherence and colonization. We have demonstrated here that cholesterol supports both increased display of Lewis X and Y antigens as well as the modification of LPS core/lipid A structure. These responses do not require cholesterol α-glycosides, but are nevertheless highly specific for cholesterol. No changes in Lewis antigen levels or in LPS profiles occurred when cholesterol was substituted by the structurally very similar β-sitosterol or other steroidal substances. There is experimental evidence for specific, protein-mediated cholesterol uptake by H. pylori , but no receptor has so far been identified.
In the clinical strain G27, specific LPS bands are observed under conditions of cholesterol depletion that do not occur upon growth in complex or defined media containing cholesterol. This suggests a requirement for cholesterol in the normal maturation of structure during LPS biosynthesis. Determination of the structure of LPS in G27, and identification of cholesterol-dependent changes to this structure, are currently in progress. We anticipate that cholesterol-dependent changes will likely be found within the core/lipid A portion of the LPS, because we also observed LPS band changes in isogenic strains that lack the O-chain. The loss of LPS O-chains by disruption of pmi was unexpected, as an NCTC11637 strain with a disruption in the same gene retained the O-chain . We do not presently know why the LPS phenotype of the latter mutant differs from the pmi::cat strains that we generated using an allelic replacement strategy. Investigation of this matter is ongoing and will be the subject of another report. Directing our attention to the core/lipid A moieties, we attempted to identify LPS biosynthesis genes that, when disabled, would eliminate the observed LPS responses to cholesterol. We selected two genes, lpxE and eptA, that sequentially remove the lipid A 1-phosphate group and add 1-phosphoethanolamine . Disruption of eptA did not affect cholesterol-dependent changes in the LPS profile, but disruption of lpxE eliminated this response to cholesterol. We propose that the LPS bands seen only under conditions of cholesterol depletion represent LPS with modified lipid A structure. This modified form could be 1-dephospholipid A, or a downstream form thereof (not including the 1-phosphoethanolamine form, which is ruled out by our eptA::cat results). While the entire sequence of LPS biogenesis has not been worked out in H. pylori, a ketodeoxyoctulosonic acid (Kdo) hydrolase activity has been detected in membrane fractions of H. pylori that removes the outermost of two Kdo residues subsequent to lipid A dephosphorylation . Though to date no Kdo hydrolase gene has been identified, such a Kdo-modified derivative may be considered a candidate for the modified LPS. There may be other as yet unidentified downstream modifications as well. Positive assignment of the bands we observed is further complicated by the existence of a minor LPS form, in which lipid A bears an extra 4-phosphate group, and is hexa- rather than tetra-acylated . Lipid A modifications are important because they strongly influence Toll-like receptor recognition, modulating innate immune responses [23, 64].
In order to discuss potential mechanisms for these LPS effects, we must consider the architecture of LPS biosynthesis. In well-studied organisms such as E. coli, the numerous steps in LPS biogenesis take place in specific subcellular compartments, and require specific transporters to shuttle intermediates across the inner membrane, periplasmic space, and outer membrane [64, 65]. Kdo2-lipid A is synthesized on the cytoplasmic face of the inner membrane, where the core oligosaccharide is separately assembled and then attached. This core-lipid A species must be flipped across the bilayer by the essential transporter MsbA. Modifications to lipid A are then carried out on the periplasmic face of the inner membrane. The O-chain is independently assembled in the cytoplasm on an undecaprenyl diphosphate carrier, transported across the inner membrane, and attached to the core-lipid A periplasmically. The multicomponent Lpt assembly transports full-length LPS across the outer membrane, where further trimming may occur. LPS biogenesis is species-specific, and for the case of H. pylori the picture is much less complete. Some but not all of the expected LPS transporter subunits have been identified in the genome [66, 67]. Lipid A dephosphorylation and phosphoethanolamine addition have been assigned to the periplasmic compartment based on work in which these H. pylori genes were expressed in a temperature-sensitive MsbA mutant strain of E. coli . Our data are consistent with periplasmic lipid A modification occurring independently of both O-chain addition and Lewis antigen addition, in keeping with the general model just described. This distinctly ordered process gives rise to a defined range of LPS molecules at the cell surface. Importantly, the LPS array can be remodeled in response to environmental conditions such as external pH [68, 69].
How then might cholesterol modulate LPS biogenesis and modification? The lipid compositions of the inner and outer membranes of gram negative bacteria are specific and distinct , but little is known about the subcellular compartmentation of cholesterol in H. pylori or other prokaryotes. We propose that the presence of cholesterol is needed to establish the proper membrane composition and structure that permit the orderly building of nascent LPS as it transits across the inner membrane/periplasmic/outer membrane compartments. In this model, altered membrane composition may influence the activity of LPS biosynthetic enzymes embedded in the membrane, leading to improper LPS modification. Alternatively, cholesterol depletion may result in dysregulation of LPS transporter function due to alterations in membrane structure and composition. The dysregulated movement of LPS among inner membrane, periplasmic, and outer membrane compartments would then result in aberrant modifications to its structure. This scenario would be consistent with the observed discrepancy between whole cell Lewis antigen levels measured by immunoblot and cell surface levels measured by ELISA. That is, it is possible that under cholesterol-depletion the Lewis antigen-bearing LPS may be less effectively transported to the cell surface. Preliminary evidence indicates that membrane cholesterol may also influence certain ABC transporters and the ComB DNA transporter in H. pylori (Hildebrandt, Trainor and McGee, unpublished results). Thus, cholesterol may support a wider range of physiological processes in the bacterial membrane than is currently appreciated.
We have demonstrated for the first time that cholesterol, though nonessential to growth of H. pylori, is nevertheless essential for gastric colonization in an animal model. We have further shown that cholesterol plays important roles in determining LPS structure as well as Lewis antigen expression, and that these biological effects are highly specific for cholesterol. LPS profiles of mutant strains lacking the O-chain retain responses to cholesterol availability, providing evidence for structural changes to the oligosaccharide core/lipid A moieties. Disruption of the lipid A 1-phosphatase gene, lpxE, eliminated the effect of cholesterol on LPS profiles, suggesting that aberrant forms of LPS that appear upon cholesterol depletion are dependent upon 1-dephosphorylation of lipid A. The roles of cholesterol in LPS structural modification and in Lewis antigen expression do not require α-glucosylation of cholesterol. Thus, cholesterol imparts these benefits independently of its previously reported role in resistance to host phagocytosis and T-cell responses, which require the alpha-glycoside metabolite of cholesterol . Together these studies serve to emphasize the critical roles that cholesterol and its metabolites in the H. pylori membrane can play in host-pathogen interactions.
This work was supported by Public Health Service grant RO1CA101931 from the National Institutes of Health and by a Bridge Award from LSUHSC-S. Our colleagues Ken Peterson and Daniel Shelver took part in discussions of the work in progress. Traci Testerman shared bacterial stocks and participated in discussions. John Staczek donated laboratory supplies, and critiqued a preliminary version of this manuscript.
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