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INTRODUCTION |
The Gram-negative bacterium Helicobacter pylori is a
prevalent pathogen of humans, and chronic infection of the gastric
mucosa by the bacterium causes recurrent gastroduodenal inflammatory disease (1). H. pylori is a major cause of chronic gastritis and plays a pivotal role in the development of both gastric and duodenal ulcers (2-4). Moreover, persistent infection with this bacterium is associated with an increased risk for the development of
gastric adenocarcinoma and primary lymphoma (5, 6). H. pylori is a chronic pathogen, and the mechanisms by which this bacterium is able to persist in the stomach and resist or evade destruction by the immune system is central to its pathogenesis (1,
7).
In part, survival of H. pylori in the stomach may be
attributed to the development of specialized characteristics, including the capacity to withstand and adapt to exposure to gastric acidity. H. pylori colonizes the gastric mucus layer where the pH
gradient ranges from pH 2 on the luminal side to almost pH 7 on the
epithelial cell surface (8). The helical shape and rapid motility of
the bacterium facilitate its movement within viscous mucus, allowing the bacterium to escape extremely low pH (7, 9). Nevertheless, H. pylori cells must survive exposure to acidic pH during the early
stages of gastric infection before colonization of the gastric mucus.
Although the mucus layer provides a partial barrier to the acid
contents of the stomach, H. pylori may encounter periodic exposure to low pH depending on the location of the bacterium in the
gastric mucosa and host gastric physiology (9). In addition, H. pylori can alter the normal gastric physiology whereby acute H. pylori infection in humans is associated with transient
hypochlorhydria, possibly facilitating enhanced intragastric survival
early in infection, whereas chronic H. pylori infection
leads to increased gastric acid secretion (10, 11). Below pH 4 survival
of H. pylori is dependent on urease (12), an enzyme whose
activity is essential for colonization (13), which liberates
NH3 from urea that has been deduced to contribute to
neutralization of gastric acidity (7). However, above pH 4 urease-independent mechanisms are involved in survival but remain to be
elucidated fully.
Like the outer membrane of other Gram-negative bacteria, that of
H. pylori contains lipopolysaccharide
(LPS)1 (14). Fresh clinical
isolates of H. pylori produce high molecular mass
smooth-form LPS, which consists of a polysaccharide (PS) O-chain, a
core oligosaccharide (OS), and a lipid moiety, termed lipid A (15, 16).
The first detailed structural analysis of H. pylori LPS
showed that the PS region of the H. pylori type strain
(NCTC 11637) was composed of an elongated, partially fucosylated N-acetyllactosamine (LacNAc) polysaccharide attached to the
core OS and terminated at the non-reducing end by mono-, di-, or
trimeric Lewisx
(Lex)2 (17).
Subsequent structural studies (18-25) and serological investigations
(26-30) have shown that the O-chains of H. pylori strains
express partially fucosylated, glucosylated, or galactosylated LacNAc
chains of various lengths that may or may not be terminated at the
nonreducing end by Lex and or Ley type 2 units,
in mimicry of normal human cell-surface glycoconjugates and of glycan
antigens found in adenocarcinoma tumors (31, 32). In addition, certain
H. pylori strains express fucosylated LacNAc chains
terminated with Lea and Leb and blood group A
antigenic determinants (22, 25, 33). The pathogenic relevance of
Lex and Ley mimicry for H. pylori
remains unclear but has been suggested to aid colonization by
camouflaging the bacterium in the gastric mucosa and by aiding
bacterial adhesion (14, 34-36), whereas in chronic infection this
mimicry may induce antibodies contributing to the development of
gastritis and influence the inflammatory response in the gastric mucosa
(7, 14, 27, 29, 30).
Cultivation of H. pylori on solid agar media in
vitro can result in a shift to production of low molecular mass
rough-form LPS lacking O-chain expression and
Lex/Ley mimicry (16, 17), but this can be
reversed and production of smooth-form LPS stabilized when strains are
grown in liquid media (16, 37). In addition to this variation,
serological investigations suggest that phase variation (also called
antigenic variation) in the type of Lewis antigen expressed on H. pylori can occur in vitro and in vivo (38,
39), but this requires verification by chemical studies because
serological investigations of Le expression in H. pylori LPS
can be misleading (22, 40). Moreover, compared with growth at neutral
pH, growth of H. pylori at low pH on solid media has been
reported to induce changes in colony morphology, cellular lipids, and
virulence properties, possibly reflecting changes in bacterial cell
wall characteristics in a low pH environment (41).
Despite these observed changes in total cellular lipids, no data are
available on the influence of low pH on H. pylori LPS chemical structure or whether such conditions could induce phase variation in Lex and Ley expression. Because
different Lewis antigens are expressed at different sites within the
gastric mucosa (31, 42, 43), the ability of the bacterium to vary
Lex and Ley antigen expression in response to
environmental conditions such as pH could aid colonization at the
different sites. In this paper, we investigated acid-induced changes in
the structure and composition of LPS and cellular lipids. Bacteria were
grown at pH 5 and 7 in a liquid medium, rather than on a solid medium,
to avoid other changes such as smooth- to rough-form LPS shift, which
may occur independently of pH (15, 16). Acid-induced changes in O-chain structure including phase variation in Lex and
Ley expression were demonstrated independent of lipid A
structure and cellular lipid composition. In addition to influencing
the type of camouflage employed by H. pylori, such changes
in the major glycolipid of the outer membrane would have an important influence on the properties of this membrane and would contribute to
the adaptation of H. pylori to its ecological niche.
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EXPERIMENTAL PROCEDURES |
Bacterial Strain and Growth Conditions--
H. pylori
strain 26695, whose complete genome sequence has been determined, was
originally isolated from a patient with gastritis (40). This strain was
grown routinely on blood agar under microaerobic conditions at 37 °C
for 48 h as described previously (15, 37). Stock cultures were
maintained at 
70 °C in trypticase soy broth containing 15% (v/v)
glycerol (45). For induction of antigenic variation, H. pylori 26695 was grown in brain heart infusion containing 2%
(v/v) fetal calf serum supplemented with 50 mM potassium
phosphate buffer at pH 7 or in the same medium at pH 5 after adjustment of the pH by the addition of HCl (37, 41). Bacteria were harvested by
centrifugation (5000 × g, 4 °C, 30 min) and washed
twice, and the pellets were lyophilized.
Isolation and Degradation of Lipopolysaccharides--
After
pretreatment of bacterial biomass with Pronase (Calbiochem), LPSs were
extracted by the hot phenol-water technique (15). The water-soluble LPS
preparations were purified by treatment with RNase A, DNase II, and
proteinase K (Sigma) as described previously (15, 45) and subsequently
by gel-permeation chromatography on a column of Bio-Gel P2 (1 m × 1 cm) with water as the eluent (22). Only one carbohydrate-positive
fraction (46) was obtained, which eluted in the high molecular mass
range, consistent with previous observations (18, 22). These intact
LPSs were used for chemical and serological analyses.
The LPSs were degraded with 0.1 M sodium acetate buffer, pH
4.2, at 100 °C for 4 h to cleave the acid-labile ketosidic
linkage between the core OS and lipid A. The water-insoluble lipid A
was removed by centrifugation (5000 × g, 4 °C, 30 min), washed, and lyophilized separately. The supernatant was
fractionated by gel-permeation chromatography on a column of Sephadex
G-50 (70 × 2.6 cm; Amersham Biosciences, Inc.) using 0.05 M pyridinium acetate, pH 4.5, as eluent and monitored with
a Waters differential refractometer. The resultant water-soluble
carbohydrate-containing fractions (46) of O-chain and core OS were
collected and lyophilized. Preparative defucosylation and
dephosphorylation of PS and core OS were achieved by treatment with
aqueous 48% hydrofluoric acid (4 °C, 16 h) as described
previously (15), neutralization with cold 25% (v/v) ammonia, and
desalting by gel-permeation chromatography on a column of Sephadex G-50
(70 × 2.6 cm). Preparation of the lipid A backbone was
accomplished by the degradative procedure described previously (45).
Briefly, after treatment with 0.1 M HCl (100 °C, 30 min), lipid A was reduced with NaBH4 and subjected to
hydrazinolysis (100 °C, 48 h) and N-acetylation with
acetic anhydride/NaOH (47), and subsequently the product was purified by gel-permeation chromatography on columns of Sephadex G-25 (50 × 2 cm) and TSK-HW40S (24 × 1 cm; Merck) (45).
Electrophoretic and Serological Analyses--
For analysis of
the macromolecular heterogeneity of H. pylori LPS by gel
electrophoresis, proteinase K-treated whole-cell lysates were prepared
as described (48). These lysates and isolated LPS were analyzed by
SDS-PAGE using a stacking gel of 5% (w/v) acrylamide and a separating
gel of 15% (w/v) acrylamide containing 3.2 M urea (15).
After electrophoresis with a constant current of 35 mA for 1 h,
the gels were fixed, and LPS was detected by silver staining (49).
Alternatively, the fractionated LPS was electroblotted onto
nitrocellulose membranes by using the buffer system of Towbin et
al. (50). Nitrocellulose membranes with transferred LPSs were
probed with mouse IgM monoclonal antibodies against Le antigens
anti-Lex (clone P12), anti-sialyl-Lex (clone
CSLEX1), anti-Ley (clone F3), anti-Lea (clone
T174), anti-Leb (clone T128) or H type I antigen
(clone 17-206) (Signet Laboratories, Dedham, MA) or against blood
group determinants anti-A, -B, or -AB (Immunocor, Norcos, GA) diluted
1:1000 as primary antibody and peroxidase-conjugated goat anti-mouse
IgM (Sigma) diluted 1:1000 as the secondary antibody as described
previously (29). Reactions in Western blots were visualized with the
Bio-Rad premixed enzyme substrate kit (2.5 ml of 4-chloro-1-naphtol in
diethylene glycol, 25 ml of Tris-buffered saline, and 15 µl of
H2O2) according to the manufacturer's instructions.
Also, an enzyme-linked immunosorbent assay (ELISA) with bacterial whole
cells was used as described previously (29) to examine the reaction of
the anti-Le and blood group antibodies with H. pylori 26695 grown at pH 7 and 5. Protein concentrations of bacterial suspensions
were determined using a commercial assay (Pierce). Subsequently,
flat-bottomed microtiter plates were coated overnight with 100 µl of
cell suspensions, with a protein concentration of 60 µg/ml, in 0.05 M NaHCO3 coating buffer, pH 9.6, and blocked with 3% (w/v) bovine serum albumin at room temperature for 2 h. The ELISA assay was performed by the procedure of Wirth et
al. (28). As described previously (29), the specificities of the antibodies in the assay were validated by their ability to bind the
respective antigen from a panel of synthetic Le and blood group
antigens (Isosep AB, Tullinge, Sweden and Dextra Laboratories, Reading, UK) and the LPSs of other H. pylori strains, NCTC
11637, P466, and MO19, of known structure (17-19). The criterion that an absorbance value of <0.1 units was considered a negative result, whereas higher values were considered positive (28), was used in these
whole-cell ELISA studies. All assays were repeated in triplicate.
Preparation, Identification, and Fractionation of
Cellular Lipids--
Lipids were extracted from bacterial biomass
using the method of Bligh and Dyer (51). Thin-layer chromatography
(TLC) of total lipids was performed on silica gel 60 plates (Merck)
using chloroform/methanol/water (75:22:3, by volume) as the solvent system. Detection was by charring with 1% (w/v) CeSO4 in
10% (v/v) H2SO4 or by staining with the
respective reagents: molybdate stain for phospholipids, ninhydrin for
amino lipids, 
-naphthol stain for glycolipids, and sulfuric
acid/acetic acid reagent for sterols and sterol esters (52). The lipids
L--phosphatidylethanolamine, L--lysophosphatidylethanolamine,
L--phosphatidyl-DL-glycerol, cardiolipin,
phosphatidylcholine, and
L--phosphatidyl-L-serine were obtained from
Sigma and used as standards on TLC plates. The relative abundance of
lipids was determined by scanning laser densitometry after charring of
TLC plates (53).
Total lipid extracts were subjected to lipid anion-exchange
chromatography on 1 ml (100 mg) Superclean LC-NH2 solid
phase extraction columns (Supelco, Bellefonte, PA) as described
previously (41, 54). Fractions were sequentially eluted with the
solvents: chloroform/2-propanol (2:1, v/v), diethyl ether/acetic acid
(98:2, v/v), acetonitrile/2-propanol (2:1, v/v), methanol, and
2-propanol/3 M methanolic HCl (4:1, v/v) to yield five
fractions, A to E (41). The two predominant phospholipids,
phosphatidylethanolamine and lysophosphatidylethanolamine, were
purified from fraction D on silica gel 60 (40-60 µm, Merck) using
chloroform/methanol/water (75:22:3, by volume) as the eluent, and the
appropriate fractions were collected and dried under N2.
The identities of lysophosphatidylethanolamine and
phosphatidylethanolamine were confirmed after acidic liberation of
fatty acids (4 M HCl, 100 °C, 4 h) and subsequent
analysis by TLC (41) and by comparison with authentic
lysophosphatidylethanolamine and phosphatidylethanolamine in
31P NMR spectroscopy (55).
Sugar Composition and Methylation Linkage Analyses--
Sugar
composition analysis of LPSs was performed by the alditol acetate
method (56). Hydrolysis of glycosidic bonds was achieved with 2 M trifluoroacetic acid at 120 °C for 2 h and
followed by reduction with NaBD4, and acetylation with
acetic anhydride (100 °C, 30 min). Alditol acetate derivatives were
identified by gas-liquid chromatography (GLC) using a Hewlett-Packard
5880 chromatograph (Avondale, PA) equipped with a DB-5 fused-silica capillary column (30 m × 0.25 mm) and a temperature program of 160 °C (1 min) to 260 °C at 3 °C/min and by GLC-mass
spectrometry (MS) on a Hewlett-Packard 5890 chromatograph equipped with
a NERMAG R10-10L mass spectrometer using the same conditions.
Enantiomeric configurations of the respective sugars were determined by
GLC analysis of the acetylated 2-(+)-butyl glycosides (for GlcN) and 2-(+)-octyl glycosides (for Gal and Fuc) by the published methods (57,
58), which were modified as described (59). Linkage analysis was
performed by methylation with CH3I in Me2SO in
the presence of sodium methylsulfinylmethanide (60), and hydrolysis was
performed as in sugar analysis. The partially methylated
monosaccharides were reduced with NaBD4 and subsequently
converted to alditol acetates. Characterization of the permethylated
alditol acetates was performed by GLC-MS using the above conditions,
and identification was performed using published data (61, 62).
Analysis of Fatty Acids--
Total fatty acids were liberated
from LPS and lipid A preparations by combined acid (4 M
HCl, 100 °C, 5 h)- and base (0.5 M NaOH, 100 °C,
1 h)-catalyzed hydrolyses and with heptadecanoic acid (C17.0) as
an internal standard and were carboxymethylated with diazomethane (15,
47). Ester-bound fatty acids were selectively liberated from
vacuum-dried lipid A by alkaline trans-esterification with sodium
methylate (0.25 M, 37 °C, 15 h) as described
previously (63). Amide-bound acyloxyacyl residues were investigated
according to the procedure of Wollenweber et al. (64).
Purified phospholipids were subjected to acid- and base-catalyzed
hydrolysis (15), and after acidification, free acids were extracted
with hexane and subsequently carboxymethylated with diazomethane. The
resulting fatty acid methyl esters from these procedures were
determined quantitatively by GLC on an HP-5 fused-silica capillary
column (Hewlett-Packard) using a temperature program of 150 °C (3 min) to 300 °C at 3 °C/min, and their identities were confirmed
by GLC-MS.
NMR Spectroscopy--
Samples were exchanged twice with
D2O. 1H NMR spectra of D2O
solutions were run on a JEOL EX-270 instrument at 75 °C or on a
Varian Inova 600 instrument at 25 °C. Two-dimensional
1H,1H DQF-COSY and
1H,13C heteronuclear single-quantum coherence
experiments were performed on the Inova 600 instrument. Chemical shifts
are reported in ppm using internal sodium
3-trimethylsilylpropanoate-d4 (
H
0.00) or internal dioxane (C 67.40). For lipid analysis,
31P NMR spectra were recorded on a Varian 500 Unity
instrument at 35 °C, spectra were broad-band
1H-decoupled, and samples were referenced to an 80% (w/v)
solution of phosphoric acid (0.00 ppm) as an external standard (45,
55).
Electrospray ionization (ESI) and Laser Desorption (LD) Mass
Spectrometry--
ESI MS was performed in the negative mode using a VG
Quattro triple quadrupole mass spectrometer (Micromass, Altrincham,
Cheshire, UK) with acetonitrile as the mobile phase at a flow rate of
10 µl/min; the samples were dissolved in aqueous 50% acetonitrile at
a concentration of ~50 pmol/µl, and 10 µl was injected via a
syringe pump into the electrospray source. LD MS was carried out with a
laser microprobe mass analyzer (LAMMA 500, Leybold AG, Cologne,
Germany). Free dephosphorylated lipid A was mixed with either NaI
or CsI to obtain fragments as well as molecular ions in the positive
ion mode as described (45, 65).
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RESULTS |
SDS-PAGE and Serological Analyses--
When analyzed in SDS-PAGE
with silver staining (Fig. 1), isolated
LPSs from H. pylori 26695 biomass grown at pH 5 and 7, LPS1 and LPS2, respectively, showed profiles characteristic of slow migrating, high molecular mass LPS with PS O-chains as reported previously (15, 16). Like the purified preparations of LPS1 and LPS2,
proteinase K-treated whole-cell lysates of biomass grown under the
differing pH conditions gave identical electrophoretic profiles,
indicating that the LPS extraction and purification procedure did not
affect the macromolecular nature of the preparations. Immunoblotting
experiments showed that the O-PS region of LPS1 expressed both
Lex and Ley, whereas LPS2 expressed only
Lex (Fig. 2). No reaction of
these LPSs with monoclonal antibodies against other Le antigens
(anti-Lea, -Leb, or -H type 1) or against blood
group determinants (anti-A, -B, or -AB) was observed. In the case of
chemical modification or selection of a particular LPS molecular
species during extraction, the same panel of antibodies was tested for
reaction in an ELISA with whole cells of H. pylori grown at
pH 5 and 7, respectively (Fig. 3). Using
the criteria previously established for the ELISA (28), the observed
expression of Lex and Ley at pH 5 and only
Lex at pH 7 was consistent with the LPS1 and LPS2
immunoblotting results, respectively.
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Fig. 1.
Silver-stained SDS-PAGE gels of LPSs of
H. pylori 26695 grown at pH 5 and 7. A, purified LPSs of H. pylori 26695 grown at pH 5 (LPS1) and pH 7 (LPS2). B, proteinase K-treated whole-cell
lysates from biomass grown at pH 5 and 7. Purified LPSs, LPS1 and LPS2,
were obtained by hot phenol-water extraction of biomass grown at pH 5 and 7, respectively, and purified by enzymatic treatments with RNase A,
DNase II, and proteinase K and subsequent gel-permeation
chromatography, whereas mini-extracts of LPSs were prepared by lysing
whole bacterial cells and subsequent treatment with proteinase K as
described under "Experimental Procedures." SDS-PAGE was performed
using a stacking gel of 5% (w/v) acrylamide and a separating gel of
15% (w/v) acrylamide containing 3.2 M urea under a
constant current of 35 mA for 1 h, after which the gels were fixed
and silver-stained (49). The silver staining procedure for LPS differs
to that for conventional protein staining in the oxidation and
development conditions used (48, 49), and under these conditions silver
has been shown to specifically bind LPS because of the high affinity
ligands present in periodate-oxidized LPS for silver compared with
proteins (48). Identical electrophoretic profiles of purified LPS and
proteinase K-treated bacterial extracts at the respective pH values
were observed.
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Fig. 2.
Immunoblots of LPSs from H. pylori
26695 grown at pH 5 (LPS1) and pH 7 (LPS2) with
anti-Lex and anti-Ley monoclonal
antibodies. Lane A, pH 5; lane B, pH 7. LPS1
and LPS 2 were prepared as described in Fig. 1 and under
"Experimental Procedures." SDS-PAGE was performed using a stacking
gel of 5% (w/v) acrylamide and a separating gel of 15% (w/v)
acrylamide containing 3.2 M urea under a constant current
of 35 mA for 1 h. The fractionated LPSs were electroblotted onto
nitrocellulose membranes (50), and the transferred LPSs were probed
with anti-Lex and anti-Ley mouse IgM monoclonal
antibodies diluted 1:1000 as primary antibody and peroxidase-conjugated
goat anti-mouse IgM diluted 1:1000 as the secondary antibody as
described under "Experimental Procedures." Positive reactions can
be seen between LPS1 and anti-Lex and -Ley
monoconal antibodies, whereas only anti-Lex monoclonal
antibody reacted with LPS2.
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Fig. 3.
Whole-cell ELISA of H. pylori
26695 grown at pH 5 (LPS1) and pH 7 (LPS2) with
anti-Lex, anti-Ley, anti-Le antigen, and
anti-blood group monoclonal antibodies. The ELISA with bacterial
whole cells was used as described previously (28,29). These data
indicate that H. pylori 26695, when grown at pH 5 expresses
Lex and Ley antigens, whereas at pH 7, only
Lex antigen is expressed. Only absorbance values >0.1
units were considered positive (28).
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Structural Analyses of the LPS Core--
Sugar analysis of the
core OS derived by mild degradation at pH 4.2 of H. pylori
LPS1 (OS1) showed the presence of fucose, glucose, galactose,
2-amino-2-deoxy-D-glucose,
D-glycero-D-manno-heptose (DD-Hep), and
L-glycero-D-manno-heptose
(LD-Hep) in the molar ratios 0.65:1.3:1.0:0.55:1.1:0.65, respectively,
together with a small amount of ribose. After dephosphorylation with
aqueous 48% hydrofluoric acid the content of LD-Hep increased to
almost twice its value, and hence, a proportion of these residues was phosphorylated.
Methylation (Table I), ESI MS, and
1H NMR spectroscopic studies of OS1 before and after
dephosphorylation indicated the same basal structure 1 (see
Structures 1 and 2
) as has been
established for the core region of H. pylori NCTC 11637 LPS
(18), except that a 2-aminoethyl phosphate (PEtn) group was
identified in contrast to a phosphate monoester group that has been
reported earlier (18-20). Thus, ESI MS (Fig.
4) revealed a doubly charged
pseudomolecular ion, [M-2H]2
, at
m/z 891.80 for 1 with a calculated
molecular mass of 1785.53 (in contrast, a peak at
m/z 870.2 would be observed for the phosphate
monoester derivative that would have a calculated molecular mass of
1742.46 Da). The 3-deoxy-D-manno-octulosonic acid (Kdo) residue at the reducing end of 1 and the other core OS was found to exist in an anhydroform (anhKdo).
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Table I
Methylation analyses of sugar linkages in core oligosaccharide from H. pylori 26695 LPS1 (OS1), O-chain polysaccharides from LPS1 and LPS2
(PS1 and PS2, respectively), and modified PS1 after treatment with
aqueous 48% hydrofluoric acid (PS1M)
The GLC retention times of the corresponding alditol acetates are
referred to 2,3,4-tri-O-methylfucose (2,3,4-Me3-Fuc,
1.00) and glucitol hexaacetate (3.39).
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Fig. 4.
The region of doubly charged pseudomolecular
ions of the ESI MS spectrum of the core OS from LPS1 (OS1).
Structure 1 corresponds to A in the spectrum at
m/z 972.7. The major compound in the mixture thus
corresponds to 1 with one additional hexose residue
(A+Hex). Heterogeneity was found to be associated with the
lack of one or two hexose residues (A-Hex and
A-2Hex, m/z 810.53 and 730.01, respectively) and attachment of up to three additional hexose residues
(m/z 1053.91, and 1134.55) or two hexose and one
pentose (ribose) residue (m/z 1120.01).
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Heterogeneity of OS1 was revealed by ESI MS (Fig. 4) and found to be
associated with the lack of one or two hexose residues (peaks of doubly
charged ions, [M-2H]2, at m/z
810.53 and 730.01, respectively) and attachment of up to three
additional hexose residues (m/z 972.7, 1053.91, and 1134.55) or two hexose and one pentose (ribose) residue
(m/z 1120.01). The major compound in the mixture
corresponded to 1 with one additional hexose residue
attached. As indicated by the methylation analysis data (Table I), this
and other additional monosaccharides may substitute the terminal Glc
residue at position 3 or 6 or position 2 of the 7-substituted DD-Hep
residue in 1 (see Refs. 18-20). The exact sites of their
attachment, as well as their sequence and anomeric configurations
remain to be determined.
The disaccharide fragment
-L-Fucp-(1
3)-
-D-GlcpNAc
present in OS1 is also a constituent of the PS O-chain (see Structures T3 and T4). In the methylation analysis of OS1, the Fuc residue appeared as 3-substituted (Table I), whereas this residue is terminal,
as followed from other data for OS1, including ESI MS data (Fig. 4).
This phenomenon has been observed previously (18) and has been
suggested to originate from incomplete methylation.
Similar analyses of OS2, the core OS derived from H. pylori
LPS2, showed essentially the same structure but with a lower degree of
chain elongation such that the contribution of compounds with more than
one additional hexose residue was negligible. The major compound in the
mixture was the Glc-lacking oligosaccharide 2 (see
Structures 1 and 2). Therefore, except for the degree of glucosylation,
there is no significant difference between the core regions of the two LPSs.
Elucidation of the Structure of the PS O-Chain from H. pylori LPS1
(PS1)--
Sugar analysis of PS1, including determination of the
absolute configurations of the monosaccharides, revealed
L-fucose, D-galactose, and
2-amino-2-deoxy-D-glucose in the molar ratios 0.4:1.4:1.0 as the main components. In addition, ribose, glucose, DD-Hep, and
LD-Hep were identified as core constituents of H. pylori LPS as described above.
Methylation analysis of PS1 (Table I) resulted in identification of
partially methylated alditol acetates derived from the major
components: terminal Fuc, terminal and 3-substituted Gal, 4-substituted
GlcNAc, and 3,4- and 4,6-disubstituted GlcNAc. These data suggested
that PS1 is branched with two different lateral sugar residues (Gal and
Fuc) and two different GlcNAc residues as branch point residues.
Comparison with the methylation analysis data for LPS1 (Table I) showed
that no terminal fucose was cleaved during mild acid degradation. In
addition, a number of minor partially methylated alditol acetates were
detected. Most of them were derived from the core region of LPS, as
determined by comparison with the methylation analysis data of OS1
(Table I). However, two minor products originated from PS1. One was
derived from 3,4,6-trisubstituted GlcNAc and showed that a minor
portion of GlcNAc residues carried two side chains. The other was from
2-substituted Gal, which is located in the terminal nonreducing LacNAc
unit of PS1 (see Structure T3).
In the 1H NMR spectrum of the PS1, a signal from
PEtn was observed at 3.28 (t, J 5.0 Hz,
CH2N) with a cross-peak to 4.13 (CH2OP) in the two-dimensional
1H,1H DQF-COSY spectrum. In the two-dimensional
1H,13C heteronuclear single-quantum coherence
spectrum of the PS1, a correlation was observed from a 1H
NMR signal at 3.28 to a 13C NMR signal at 41.9, further supporting the presence of a PEtn group.
Treatment of PS1 with aqueous 48% hydrofluoric acid resulted in a
modified polysaccharide (PS1M) that eluted from Sephadex G-50 with a
similar elution volume as PS1 soon after the void volume of the column.
Sugar analysis of PS1M revealed Gal and GlcNAc in the same ratio as in
PS1 but showed only a trace amount of Fuc, indicating essentially
complete defucosylation of the polysaccharide. The content of the core
OS constituents was negligible as well, thus demonstrating cleavage
between the PS and the core region of LPS. In fact, two core-related
oligosaccharides were isolated after the reaction, one eluted with the
same elution volume as the OS1, i.e. approximately twice the
void volume, and the other just after.
Methylation analysis of PS1M (Table I) yielded terminal and
3-substituted Gal and 4- and 4,6-disubstituted GlcNAc in the ratios
0.58:1.00:0.30:0.23, respectively. Therefore, in PS1M the lateral Gal
residue is attached at position 6 of GlcNAc, and hence, in PS1 the
fucose residue substitutes GlcNAc at position 3. A minority of the
GlcNAc residues carries both fucose and galactose, which is deduced
from the presence of 3,4,6-trisubstituted GlcNAc in the methylation
analysis of PS1 (Table I). As judged by the ratios of methylated
derivatives from 4-substituted, 3,4-disubstituted, 4,6-disubstituted,
and 3,4,6-trisubstituted GlcNAc, 29% of the LacNAc units in PS1 are
not substituted (structural unit
3B), 25% are fucosylated
(structural unit 3A and 3C), 42% are
galactosylated (structural unit 3D), and 4% carry both
lateral Gal and Fuc (structural unit 3E).
The 1H NMR spectrum of PS1M contained, among other things,
signals for three anomeric protons at 4.47 (-Gal H1, split to two close doublets, J1,2 7.5 Hz for each), 4.74 (-GlcNAc H1, J1,2 ~8 Hz), and 5.02 (-Gal
H1, J1,2 2.5 Hz; the coupling, lower than
expected, is probably due to second order effects). The ratios of the
intensities of the anomeric signals were 1:1:0.45, respectively, which
fitted well with the methylation analysis data. Therefore, the
1H NMR data confirmed the presence of a poly(-LacNAc)
chain and showed that the lateral Gal residue is -linked.
The 1H NMR spectrum of PS1 indicated a higher degree of
structural heterogeneity due to nonstoichiometric substitution with two
lateral monosaccharides, Gal and Fuc. The major signals for anomeric
protons belonged to -Gal and -Fuc (both at 5.04, J1,2 ~3 Hz), -GlcNAc ( 4.69, J1,2 8.6 Hz), and -Gal ( 4.46, J1,2 7.9 Hz), the assignment based on
two-dimensional 1H,1H DQF-COSY and
1H,13C heteronuclear single-quantum
coherence3 experiments. Also,
the chemical shifts of the C5 signals of the Fuc groups were observed
at ~ 67.5 ppm, corroborating the -anomeric configuration.
Furthermore, the 1H NMR spectrum of PS1 contained signals
for N-acetyl groups of GlcNAc ( 2.02) and
CH3-C groups (H6) of Fuc ( 1.14-1.26) with the ratio of
intensities 1:0.35. In total, there were H6 signals for five Fuc
residues, two of which appeared as separate doublets at 1.23 and
1.26 and three others as three superposed doublets at 1.14 (J5,6 ~6 Hz for all H6 signals, Fig.
5A). The two-dimensional DQF-COSY spectrum (Fig. 6) showed Fuc
H6/H5 cross-peaks at 1.14/4.35, 1.14/4.81, 1.23/4.87, and
1.26/4.25. The first cross-peak was from the Fuc residue, which is
adjacent to the LPS core region, as deduced from the DQF-COSY spectrum
of OS1, where only one Fuc H6/H5 cross-peak was observed, and at the
same coordinates (compare also the one-dimensional 1H NMR
data, Fig. 5, B and C). The cross-peak at 1.26/4.25 was assigned to the Fuc group substituting position 2 of
-Gal in the terminal nonreducing LacNAc unit (compare
e.g. with 4.22 for Fuc H-5 in
2'-(-fucopyranosyl)lactose (66)). The cross-peak at 1.23/4.87,
having the same intensity, originated from the Fuc residue of the
Fuc-(13)-GlcNAc unit in a terminal Ley tetrasaccharide
(24). The last cross-peak at 1.14/4.81 of double intensity was from
the Fuc group of the same Fuc-(13)-GlcNAc fragment in two of the
interior repeating units of PS1. The latter assignment was consistent
with the chemical shift ( 4.83) of the signal for Fuc H5 in a human
milk oligosaccharide (LNF-III), which has a similar fragment (67).
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Fig. 5.
Part of 1H NMR spectra of
oligosaccharides and polysaccharides derived from H. pylori
26695 showing the H-6 resonance region of the fucose
residues. A, O-chain polysaccharide from LPS1 (PS1).
B, O-chain polysaccharide from LPS2 (PS2). C,
core oligosaccharide from LPS1 (OS1). Assignments are as follows. The
peak at 1.26 was assigned to the Fuc group substituting position 2 of -Gal in the terminal nonreducing LacNAc unit. The peak at 1.23 originated from the Fuc residue of the Fuc-(13)-GlcNAc unit in
a terminal Ley tetrasaccharide. The peak at 1.17 was
from the Fuc residue of the Fuc-(13)-GlcNAc unit in a terminal
Lex trisaccharide. The signals at 1.14 originate from
Fuc in LacNAc units of the main chain and Fuc adjacent to the LPS core
region.
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Fig. 6.
Region for fucose H-5/H-6 correlation in the
two-dimensional DQF-COSY spectrum of the O-chain polysaccharide from
LPS1 (PS1). The assignments of signals are described in Fig. 5 and
in the text.
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Therefore, two Fuc residues are present at the nonreducing LacNAc unit
of PS1, where they form a Ley unit (structural unit
3A), two more are attached to two interior LacNAc units
(structural units 3C and 3E), and the fifth Fuc
residue is attached to the unit that is adjacent to the LPS core. The
structure of PS1 may be described by formula 3, in which the
exact distribution of various interior structural units B-E along the
PS chain is unknown. Based on the ratios of the GlcNAc
derivatives revealed in methylation analysis (Table I) and on the Fuc
and GlcNAc methyl group signal intensities in the 1H NMR
spectrum, the average degree of polymerization in PS1 was estimated as
12-13 LacNAc units.
Elucidation of the Structure of the PS O-chain from H. pylori LPS2
(PS2)--
Sugar analysis showed that the content of Gal in PS2 was
significantly lower than in PS1 (the ratios of Fuc:Gal:GlcN were 0.35:0.95:1). Methylation analysis revealed the absence of
4,6-disubstituted GlcNAc and the presence of only a small amount of
terminal Gal, whereas the content of the other PS constituents
(terminal Fuc, 3-substituted Gal, 4-substituted GlcNAc, and
3,4-disubstituted GlcNAc) indicated a close similarity between PS2 and
PS1 (Table I). As deduced from the ratio of methylated GlcNAc
derivatives, the degree of fucosylation in PS2 was 30%. No
2-substituted Gal was detected, but instead, terminal Gal was present
(Table I), and therefore, unlike PS1, PS2 has no Ley
antigenic determinant.
The 1H NMR spectrum of PS2 was similar to that of PS1 but
lacked the H1 signal for -Gal. Also, the signals for H6 of Fuc
displayed a different pattern; there was a doublet at 1.17 and
superposition of three doublets at ~ 1.14 (J5,6 ~6 Hz for all H6 signals, Fig.
5B). As expected, signals at 1.23 and 1.26, which
belonged to the Fuc residues from the terminal Ley
tetrasaccharide in PS1 (Fig. 6), were absent from the spectrum of PS2.
The ratio of the integral intensities of the signals for methyl groups
of Fuc ( 1.14-1.17) and GlcNAc ( 2.03) was 0.32:1, again
indicating incomplete fucosylation of LacNAc units.
The two-dimensional DQF-COSY spectrum of PS2 showed, among other
things, Fuc H6/H5 cross-peaks at 1.14/4.33, 1.14/4.81, and
1.17/4.83. The two first cross-peaks were also present in the spectrum
of PS1 (Fig. 6) and belonged to Fuc residues attached to the core
region and to the interior LacNAc units, respectively (see Structure
T3). The last cross-peak was absent from the spectrum of PS1 and
was assigned to the Fuc residue located at the terminal nonreducing
LacNAc unit of PS2, where it forms an Lex unit (24). As in
PS1, there are only two Fuc residues attached to interior LacNAc units,
and the average degree of polymerization of PS2 was estimated as 12 LacNAc units.
These data suggest that PS2 has structure
4, which contains terminal fucosylated LacNAc (structural
unit 4A) and both non-fucosylated (structural unit
4B) and fucosylated LacNAc (structural unit 4C).
This structure differs from structure 3 of PS1 only in the
absence of substitution by lateral -Gal residues and the second
-Fuc residue from the nonreducing LacNAc unit, producing a terminal
Lex trisaccharide (structural unit 4A) rather
than Ley tetrasaccharide.
Structural Analysis of Lipid A--
Compositional analysis of free
lipid A preparations from LPS1 and LPS2 (LA1 and LA2) revealed, beside
fatty acids, a similar composition of GlcN and phosphate in the molar
ratio 2:1.4, with trace amounts of ethanolamine. The fatty acids
present in both lipid A preparations were dodecanoic (C12),
tetradecanoic (C14), hexadecanoic (C16),
octadecanoic (C18), 3-hydroxyhexadecanoic (OHC16), and 3-hydroxyoctadecanoic (OHC18)
acids in the approximate molar ratios 0.5:0.4:0.1:1.0:1.3:2.0, respectively.
Treatment of these lipid A preparations with 0.1 M HCl
(100 °C, 30 min) liberated glycosidic phosphate (1 eq), and
subsequent reduction with NaBH4 yielded products containing
D-GlcN and D-glucosaminitol (1.0:0.9) with
residual phosphate content (0.4 eq), attributable to ester-bound
phosphate as reported previously (45). Subjecting these products to the
chemical degradation pathway for lipid A backbone analysis developed
earlier (45, 47) yielded N-acetylated disaccharides of
GlcNAc1-6GlcNAc-ol with identical properties in high pressure
liquid chromatography, identical 1H NMR spectra, and after
permethylation, identical GLC-MS mass spectra as authentic standards
(47). Analysis by 31P NMR spectroscopy of both LA1 and LA2
revealed a signal (2.96 ppm) for a glycosidic phosphomonoester and a
second for a phosphodiester (1.32 ppm), which were attributed to
glycosidic phosphate and ethanolamine-phosphate groups, respectively,
since acidic treatment (0.1 M HCl, 100 °C, 30 min) led
to their liberation and loss of both signals. A further but weaker
signal corresponding to an ester-bound phosphomonoester (4.52 ppm),
which was acid-stable, was attributed to a 4'-phosphate group (45).
Collectively, these data showed that the backbones of LA1 and LA2 were
identical 1,4'-bisphosphorylated (1,6)-linked GlcN disaccharides.
To study the distribution of fatty acids on the lipid A backbone,
dephosphorylated LA1 was subjected to LD MS after cationization by a
NaI admixture and analyzed in the positive-ion mode (Table II). In addition, analysis of
dephosphorylated LA1 by LD MS after admixture of CsI gave a similar
fragmentation pattern and showed the presence of nonstoichiometric
amounts of amide-bound C16OC18 on the
nonreducing GlcN unit of the lipid A backbone (data not shown).
Collectively, the data showed that the reducing GlcN of the backbone
carries ester-bound 3-hydroxyoctadecanoic and ester-linked (OHC16), whereas the nonreducing GlcN can carry amide-bound
C18OC18 and ester-bound
C12OC16 or C14OC16
(Fig. 7), identical to the fatty acid
distribution reported previously in H. pylori NCTC 11637 (45). However, heterogeneity in the acylation pattern in lipid A from
smooth-form LPS of H. pylori has been observed whereby
tetraacyl lipid A predominates, but hexaacyl lipid A is also present
(45, 68). Therefore, to resolve this issue, dephosphorylated LA1 was
subjected to silica gel chromatography (47), and one major and a second
minor fraction were obtained. Analysis by LD MS of these fractions
showed the predominance of tetraacyl lipid A (without ester-bound fatty
acids bound to the nonreducing GlcN unit), whereas analysis of the
minor fraction showed the hexaacyl distribution (Table II). Identical
data were obtained for LA2. Collectively, these results show no
comparative difference in the structure of LA1 and LA2.
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Table II
Assignment of peaks in the LD mass spectra of dephosphorylated lipid A
derived from H. pylori 26695 LPS1 (LA1)
The relevant cleavage process and the resultant structure are shown in
Fig. 7. M indicates the quasimolecular ion derived from the acylated
GlcN disaccharide; ML and MH indicate the fragment ions
derived from the acylated reducing and nonreducing GlcN units of the
lipid A backbone.
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Fig. 7.
Interpretation of the fragmentation pattern
of dephosphorylated lipid A derived from LPS1 (LA1). The
numbers in circles refer to the number of carbon
atoms in acyl chains, and the letters indicate the
designated cleavage process. Refer to Table II for details of the
formation of positive ions.
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Composition of Cellular Lipids--
The content of total lipids in
H. pylori 26695 grown at pH 5 and 7 was similar (7.2 and
7.4% w/w, respectively). The neutral phospholipid fraction from both
contained phosphatidylethanolamine, lysophosphatidylethanolamine,
phosphatidylserine, and phosphatidylcholine at 91.6, 8.3, ~0.05, and
~0.05% (w/w), respectively. In contrast to a previous report (41),
there was no elevation in phosphatidylserine content when this H. pylori strain was grown at pH 5. Furthermore, the fatty acid
composition of phosphatidylethanolamine of H. pylori 26695 grown at pH 7 was C14 (29.5%), C15 (1.7%),
C16 (3.6%), C17 (0.3%), C18
(4.6%), C18:1 (4.6%), C18:2 (0.2%),
C19 (1.1%), C19cyc (53.4%) but did not differ
significantly from that of growth at pH 5.
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DISCUSSION |
The present investigation on the chemical composition of LPSs from
H. pylori grown at different pH values has added new insight into the expression of Lex and Ley mimicry by
this bacterium, particularly phase variation. When grown in liquid
medium at pH 7, the O-chain of H. pylori 26695 (PS2)
consisted of a LacNAc polysaccharide that was glycosylated with
-L-Fuc at O-3 of the majority of GlcNAc residues forming Lex units, including chain termination by a Lex
unit. However, growth in liquid medium at pH 5 resulted in production of a more complex O-chain (PS1) whose backbone of LacNAc units was
partially glycosylated with -L-Fuc, thus forming
Lex, whereas the majority of the nonfucosylated GlcNAc
residues were substituted at O-6 by -D-Gal residues, and
the chain was terminated by a Ley unit. Thus, the
nonreducing termini differ in the two O-chains as well as in
glycosylation of the internal LacNAc units. In contrast, detailed
chemical analysis of the core and lipid A components of LPS and
analysis of cellular lipids did not show significant differences
between H. pylori 26695 grown at pH 5 and 7. Type 2 series,
Lex, Ley, and sialyl-Lex, and in
addition, type 1 determinants, Lea, Leb, and
H-1 antigen, have been reported previously in structural studies of PS
O-chains of individual H. pylori strains (17-20, 22-25,
33). Nevertheless, in serological surveys of strains from different
geographical regions, Lex and Ley predominate,
expressed by >80% of strains (26-30). Consistent with the results of
this study, previous structural studies on other H. pylori
strains have reported that Ley, when present in smooth-form
LPS, occurs as the terminal unit on the O-chain, whereas
Lex occurs terminally and as an internal unit in the
O-chain (17-20, 22-25). However, glycosylation of internal LacNAc
units of the H. pylori O-chain with hexoses is rarer and,
although reported to occur at O-6 of nonfucosylated GlcNAc residues
previously (21, 23), can occur in O-chains not expressing
Lex or Ley (21). Thus, the observation that,
when grown at different pH values, the same H. pylori strain
can express two O-chains differing in O-fucosylation
patterns, and hence Lex and Ley mimicry, as
well as differing in occurrence of galactosylation is a novel finding.
A previous study by Bukholm et al. (41) that examined the
cellular lipid composition of H. pylori grown on solid media
at pH 5 and 7 found a much reduced amount of phosphatidylethanolamine, a predominant amount of lysophosphatidylethanolamine, and elevated phosphatidylserine at the lower pH, whereas phosphatidylethanolamine predominated, and phosphatidylserine was a minor constituent at neutral
pH. Also, the fatty acid composition of phosphatidylethanolamine differed at pH 5 and 7. The investigators suggest that the differing colony morphology and virulence properties of the strain grown at the
lower pH reflected changes in the polarity of the bacterial cell wall
due to a changed lipid composition because of a bacterial response to
the acidic environment. In contrast, in the present study no elevation
in content of lysophosphatidylethanolamine and phosphatidylserine was
observed for H. pylori 26695 grown in a liquid medium at pH
5 nor did the fatty acid composition of phosphatidylethanolamine differ
at pH 5 and 7 in liquid media. Likewise, no significant differences
were found between lipid A components of bacteria grown in liquid media
at pH 5 and 7. The observed changes in lipid composition on solid media
by Bukholm et al. (41) may be dependent on the physical
nature of the growth medium rather than pH. Consistent with this,
prolonged subculturing and growth of H. pylori on solid
medium in vitro, independent of pH, can induce changes in
cellular lipid composition4
that are associated with loss of the O-chain and a shift from smooth-
to rough-form LPS (15, 16). In contrast, fresh clinical isolates of
H. pylori produce smooth-form LPS, indicating that high
rather than low molecular mass LPS is produced in vivo (15). However, growth of H. pylori in liquid media in
vitro stabilizes production of smooth-form LPS containing a PS
O-chain (16, 37), and thus, these conditions were used in the present
study when examining the influence of environmental pH.
Further demonstrating the importance of the physical nature of the
growth medium in these investigations, H. pylori 26695, when
grown on solid medium, was shown previously to produce a low molecular
mass semi-rough LPS carrying a single Le unit, mostly a Ley
unit, but alternatively, type 1 and type 2 linear B blood group and
Lex attached to the core OS, whereas when grown in liquid
medium, the strain produced smooth-form LPS (25). The O-chain of the latter was composed of a polyfucosylated LacNAc chain terminated with a Lex unit, corresponding to PS1 in this study.
Although no significant differences were observed between the core OS
of bacteria grown in liquid media at pH 5 and 7 in the present study,
these cores differed from that of H. pylori 26695 when grown
on solid media at neutral pH (25) by the predominant absence of a
glucan chain substituting -Gal on the fourth heptose (DD-Hep) residue.
The genome of H. pylori 26695 and that of another strain,
J99, contain two copies of (1,3)-fucosyltransferase
(HpfucT) genes that are required for expression of
Lex and Ley but differ in the number of a
seven-amino acid sequence repeat, YDDLRVN (44, 69). DNA motifs near the
5'-end of these genes (HP0379 and HP0651) at two distinct
polynucleotide repeats have been deduced to indicate regulation through
slipped-strand repair (44). No putative gene for
(1,2)-fucosyltransferase (HpfucT2), which is required for
Ley synthesis, was initially identified in the 26695 genome
(44), but a truncated gene (HP0094) with a C14 tract was found, and in silico insertion of a C-G pair yielded a full-length
protein with strong homology to (1,2)-fucosyltransferase (70).
Moreover, sequence analysis has shown that the HpfucT2 gene
contains a hypermutable sequence (poly-C and TTA repeats) that provides
a possibility of frequent shifting into and out of coding frame by a
polymerase slippage mechanism (71). Phase variation in expression of
Lex has been attributed to changes in the lengths of poly-C
tracts in the HpfucT genes (38), but the length of these
tracts has not been found to be predictive of the phenotype (72). On
the other hand, variable expression of Ley by H. pylori strains has been proposed to occur at the combined levels
of replication slippage (mutation), transcription, and translation of
the fucT2 gene (71). Despite these deductions as to the
putative molecular mechanisms involved in variable Lex and
Ley expression by H. pylori, the resultant
phenotypes of phase variants were not established previously in
detailed structural studies. Furthermore, the environmental trigger
inducing variable Lex and Ley expression was
not identified.
The present study addresses such issues by showing that relative pH can
influence expression of Lex and Ley,
particularly at the termini of the O-chains, but also that pH influences glycosylation, including substitution at O-6 by
-D-Gal residues of the internal LacNAc units of these
chains. Consistent with these structural findings as well as the
serological and electrophoretic investigations in this study, McGowan
et al. (73) found qualitative differences in LPS
electrophoretic profiles when H. pylori 60190 was grown at
pH 5 compared with pH 7. Using subtractive RNA hybridization they
identified an acid-inducible gene in this strain whose protein was
highly homologous to that of WbcJ of enteric bacteria that is
considered involved in the conversion of GDP-D-mannose to
GDP-D-fucose. Moreover, a corresponding gene (HP0045)
occurs in the 26695 genome, which had previously been designated a
nolK homologue based on 44% identity with NolK, an
inducible nodulating protein of Azorhizobium caulinodans
(44). Thus, mechanisms affecting fucose availability in addition to fucosyltransferase activity are present in H. pylori 26695 that can influence Lex and Ley expression.
Common to the genomes of both sequenced strains, H. pylori
26695 and J99 are seven open reading frames encoding putative
glycosyltransferases (HP0159/JHP147, HP0208/JHP194, HP0619/JHP563,
HP0679/JHP620, HP0805/JHP741, HP0826/JHP765, and HP1105/JHP1031) that
have been implicated in LPS core synthesis, but in addition there are
three strain-specific open reading frames (JHP562, JHP820, and JHP1032)
in strain J99 and one (HP1578) in strain 26692, reflecting differences
in the core OS of the two strains (25, 69). Three genes in the 26695 genome (HP0159, HP0208, and HP1416) are homologues of the
(1,2)-glucosyltransferase gene (rfaJ) found in enteric
bacteria (69, 74), and although it has been debated whether some of
these genes encode (1,4)-galactosyltransferase and/or
(1,3)-N-acetylglucosaminyltransferase functions needed for type 2 chain synthesis (70), (1,2)-substituted Glc can occur in
the core of H. pylori 26695 under certain growth conditions (25). Also, homologues of galactosyltransferases from Klebsiella pneumoniae have been suggested to be involved in O-chain synthesis in strain 26695 (74). However, rather than involvement in addition to
(1,4)-substituted Gal in type 2 chains, these may be required for
addition at O-6 of nonfucosylated GlcNAc residues by
-D-Gal residues, as observed in PS1.
Several enteric bacteria are known to vary gene expression in response
to acid stress (75). However, there has been no evidence that exposure
to acidic pH alters expression of their LPS-associated genes. On the
other hand, compared with gonococcal cells grown at pH 8.2, those grown
at pH 5.8 produce LPS with an altered electrophoretic profile,
indicative of pH regulation (76), but this has not been characterized
structurally. In Rhizobium leguminosarum, LPS is modified in
response to several environmental stresses, including low pH (77). Loss
of a plasmid carrying acid tolerance genes from R. leguminosarum results in interference with ability to maintain
intracellular pH homeostasis at low external pH, but also ablation of
O-chain biosynthesis occurs with an acid-sensitive phenotype (78). In
contrast, as demonstrated in this study, H. pylori varies
glycosylation of the O-chain in response to acid stress, producing a
more complex structure at pH 5 than pH 7, and genes carried on the
bacterial chromosome are implicated in this phenomenon.
The in vitro observations of the present study are of
in vivo relevance since serological analyses of sequential
H. pylori isolates from patients have shown that expression
of Lex and/or Ley by isolates from the same
patient varied over time, although isolates appeared genetically
identical by analysis of randomly amplified polymorphic DNA-polymerase
chain reaction patterns (39, 79). A definite biological role for this
variation has not yet been defined, although antigenic variation in
other bacteria allows increased persistence and/or pathogenicity (80).
Potentially, the ability to vary expression of Lex and
Ley could influence bacterial camouflage in the gastric
mucosa (34), particularly since Le expression varies in different
regions of this mucosa and is dependent on the secretor status of the
individual (31, 32, 42, 43). However, the relevance of H. pylori expression of Lex and Ley for
camouflage in the gastric mucosa has been questioned (30, 81). On the
other hand, polymeric Lex expression has been implicated in
adhesion of H. pylori to human antral gastric mucosa (14,
35). Moreover, the majority of H. pylori in infected hosts
are free-living in the mucus layer, and only a proportion (about 20%)
appear to adhere to epithelial cells of the gastric mucosa (9). In the
gastric mucus layer, the pH gradient ranges from pH 2 on the luminal
side to almost pH 7 on the epithelial cell surface (8). Compared with
expression of an O-chain predominantly composed of Lex
units and terminated by a Lex unit at pH 7 (PS2) and, thus,
with optimal expression of Lex for adhesion, production of
an O-chain with lesser Lex units and capped with a terminal
Ley unit at pH 5 (PS1) can be hypothesized to affect the
availability of the Lex ligand for interaction with the
gastric mucosa. Hence, the influence of environmental pH on
Lex-mediated adhesion by H. pylori remains an
important question for further investigation.
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ABSTRACT
INTRODUCTION
