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J Biol Chem. 2008 June 6; 283(23): 16037–16050. doi: 10.1074/jbc.M709615200. | PMCID: PMC2414287 |
Copyright © 2008, The American Society for Biochemistry and
Molecular Biology, Inc. Structural Characterization of the Primary O-antigenic Polysaccharide of
the Rhizobium leguminosarum 3841 Lipopolysaccharide and
Identification of a New 3-Acetimidoylamino-3-deoxyhexuronic Acid Glycosyl
Component A UNIQUE O-METHYLATED GLYCAN OF UNIFORM SIZE, CONTAINING
6-DEOXY-3-O-METHYL-D-TALOSE, N-ACETYLQUINOVOSAMINE, AND RHIZOAMINURONIC ACID
(3-ACETIMIDOYLAMINO-3-DEOXY-D-GLUCO-HEXURONIC
ACID)*![[S with combining enclosing square]](/corehtml/pmc/pmcents/x0053x20DE.gif) L. Scott Forsberg‡ and Russell W. Carlson‡1 ‡Complex Carbohydrate Research Center, University of Georgia, Athens, Georgia 30605 Received November 26, 2007; Revised April 1, 2008. Rhizobium are Gram-negative bacteria that survive intracellularly,
within host membrane-derived plant cell compartments called symbiosomes.
Within the symbiosomes the bacteria differentiate to bacteroids, the active
form that carries out nitrogen fixation. The progression from free-living
bacteria to bacteroid is characterized by physiological and morphological
changes at the bacterial surface, a phase shift with an altered array of cell
surface glycoconjugates. Lipopolysaccharides undergo structural changes upon
differentiation from the free living to the bacteroid (intracellular) form.
The array of carbohydrate structures carried on lipopolysaccharides confer
resistance to plant defense mechanisms and may serve as signals that trigger
the plant to allow the infection to proceed. We have determined the structure
of the major O-polysaccharide (OPS) isolated from free living Rhizobium
leguminosarum 3841, a symbiont of Pisum sativum, using chemical
methods, mass spectrometry, and NMR spectroscopy analysis. The OPS is composed
of several unusual glycosyl residues, including
6-deoxy-3-O-methyl-d-talose and
2-acetamido-2deoxy-l-quinovosamine. In addition, a new glycosyl
residue, 3-acetimidoylamino-3-deoxy-d-gluco-hexuronic acid
was identified and characterized, a novel hexosaminuronic acid that does not
have an amino group at the 2-position. The OPS is composed of three to four
tetrasaccharide repeating units of
→4)-β-dGlcp3NAmA-(1→4)-[2-O-Ac-3-O-Me-α-d-6dTalp-(1→3)]-α-l-Fucp-(1→3)-α-l-QuipNAc-(1→.
The unique 3-amino hexuronate residue, rhizoaminuronic acid, is an attractive
candidate for selective inhibition of OPS synthesis. Rhizobium leguminosarum is a Gram-negative endosymbiont that forms
a nitrogen-fixing symbiosis with the legume host Pisum sativum. Like
other Rhizobiaceae, it is a member of the α-2 subgroup of the
Proteobacteria, which includes the phytopathogen Agrobacterium and
phylogenetically related bacteria such as the intracellular animal pathogens
Bartonella and Brucella
( 1,
2). A significant feature
shared by members of this subgroup is the ability to survive intracellularly
within the eukaryotic host, often surrounded by host membrane-derived
compartments, which in the case of rhizobia are termed symbiosomes. Although the early stages of symbiotic infection have been studied, factors
enabling rhizobia to survive within the host cell environment throughout their
life cycle are poorly understood
( 2- 7).
This is due in part to the difficulty in obtaining sufficient quantities of
purified bacteroid mass to allow structural study of components. A model for
symbiotic infection begins with a mutual exchange of signal molecules,
including inter alia plant flavonoids and bacterial
lipochitooligosaccharides, leading to bacterial adhesion to root hairs and the
induction of unique plant-derived structures, e.g. root nodules,
infection threads, and symbiosomes
( 2,
8). Rhizobia migrate through
the infection threads and are internalized into the root cortical cells
through a process resembling endocytosis. Internalization yields symbiosomes,
specialized intracellular compartments composed of a plant-derived membrane
that closely surrounds the bacterium. Within the symbiosomes the rhizobia
differentiate into bacteroids, the active form that reduces atmospheric
nitrogen to ammonia. In the case of R. leguminosarum, histochemical
and electron micrograph studies have shown that the rhizobial surface is in
close proximity to the surrounding plant-derived symbiosome membrane and that
contact points appear to exist between the two
( 2,
9,
10). The lipopolysaccharides
(LPS) 2 are major
structural and antigenic components of the rhizobial outer membrane, and are
suitably located to interact with the plant membrane components and soluble
plant products existing in the peribacteroid space
( 2- 5,
11,
12). Numerous studies with
rhizobial LPS mutants containing structurally defined defects have indicated
that a structurally intact LPS, expressed at normal levels, is essential for
normal root nodule development and active nitrogen fixation (Ndv +,
Fix + phenotype) ( 2,
5,
6,
13- 16). Compositional and immunological studies have shown that in R.
leguminosarum and the closely related Rhizobium etli,
LPS/O-antigen epitope expression is modified by environmental factors,
including growth at acidic pH or low oxygen concentration, conditions thought
to mimic those within the nodule
( 17- 20).
In the majority of cases the epitope structures have not been characterized,
and it has been difficult to draw precise conclusions about the significance
of structural alterations with regard to nodulation efficiency or bacteroid
survival. An advance in our understanding of rhizobial LPS structure-function
was the observation that the expression of O-antigen/LPS structure can change
dramatically upon progression from the free-living state to the bacteroid
form. The structural details of this phase shift in LPS surface chemistry were
recently described in the Sinorhizobium sp. NGR234 model system,
where LPS expression shifts from that of a structurally complex, highly
branched anionic rough LPS (lacking O-antigen) to an endogenously methylated,
hydrophobic, rhamnan O-antigen (attached to a structurally modified core lipid
A, i.e. a smooth LPS)
( 6). These structural changes
also occur on nitrogen-fixing bacteroids isolated from host Vigna
unguiculata nodules ( 6,
15). In the other model system
recently studied, that of R. etli- Phaseolus vulgaris, the
transition from free-living state to bacteroid was accompanied by specific
changes in the location and extent of O-methylation of O-antigen,
changes that could also result in an alteration of bacteroid surface
hydrophobicity ( 7). These and
related studies indicate that a variety of changes in LPS surface chemistry
occur during or immediately prior to differentiation of the free-living
rhizobia (as exists within the infection threads) into bacteroids
( 2,
3,
5,
9,
15,
21- 23),
and the changes appear to be host-symbiont-specific. The significance of these
changes is not entirely clear, but may be involved in preparing the bacterial
surface for long term survival within the symbiosome by proper interaction
with the plant membrane or attenuation of host defense mechanisms
( 3- 5,
18). The lipid A portion of R. leguminosarum 3841 LPS was examined
structurally and functionally, and an acpXL mutant was found
defective in its ability to transfer long chain fatty acid to lipid A when
cultured under normal conditions
( 24). The mutant was restored
in its synthesis of long chain fatty acidlipid A when grown in the pea nodule
environment, indicating that lipid A structure could be influenced by the host
( 24). Continuing our analysis
of the R. leguminosarum- Pisum model system, we have
characterized the structure of the O-polysaccharide portion of the LPS from
the free-living form of R. leguminosarum 3841 bv. viciae.
Like R. etli, this bacterium synthesizes a structurally complex, low
molecular mass OPS of uniform size, with endogenous O-methylation and
O-acetylation contributing to structural heterogeneity. In addition,
we have identified a new glycosyl residue,
3-acetimidoylamino-3-deoxy- d- gluco-hexuronic acid
(Glc3NAmA, rhizoaminuronic acid) as a component of the 3841 OPS repeating
unit. This is apparently the only known hexosaminuronic acid lacking an amino
group at C2. Extensive structural differences between the bacteroid and
free-living forms of the R. leguminosarum 3841 OPS exist and will be
described in a forthcoming report. The availability of the complete genome
sequence of R. leguminosarum 3841
( 25), together with structural
information, will help clarify the role of specific LPS structural features in
bacteroid adaptation and survival. Growth of Bacteria— R. leguminosarum strain 3841 was
grown in liquid fermentor culture at 28 °C in tryptone/yeast extract (TY)
supplemented with Ca 2+ as described for related rhizobia
( 12). Cells were pelleted at
late log phase ( A600 = 2.50) and then washed by
resuspending in phosphate-buffered saline followed by centrifugation to remove
exopolysaccharides and culture media, yielding 648 g (wet weight) of cells per
100 liters of culture. Isolation of Lipopolysaccharide and O-polysaccharide—The
washed cells were extracted using a hot phenol/water procedure in which the
water layer contained 5 m m EDTA, 0.05% sodium azide, and 50
m m Na 2HPO 4 buffer, pH 7.0, as described
previously ( 11).
Lipopolysaccharides (LPSs) were isolated from the resulting water layer using
standard chromatographic procedures
( 21,
26) (details are described in
supplemental Fig. S1). The total LPS was subjected to mild hydrolysis in 10
m m sodium acetate buffer, pH 4.5, for 4 h at 105 °C. Lipid A
was removed by ultracentrifugation, and the polymeric carbohydrate portion
(O-polysaccharide; OPS) was isolated by chromatography using a Bio-Gel P-10
column (45-90 μm, 1.5 × 90 cm). Further procedures describing lipid A
and core oligosaccharide removal are in the supplemental material. Fractionation of O-polysaccharides by HPLC—The void volume
from the Bio-Gel P-10 column, which contained the total soluble OPSs, was
dialyzed, concentrated, and fractionated into separate polysaccharide
components using an Asahipak-NH2P-50 4E column (4.6 × 250 mm, 5 μm)
equipped with matching guard column (4.6 × 10 mm) and NH2P-LF
pre-injector line filter (Shodex, Showa Denko K.K., Tokyo, Japan, distributed
by Thomson Instrument Co.). The column was eluted at 0.7 ml/min at 35 °C,
with a linear A/B gradient starting at 5% reagent B and ending at 70% B over
~1 h. Reagent A consisted of 75% acetonitrile, 25% water, v/v, and reagent
B consisted of 20% acetonitrile, 80% aqueous 1 m ammonium formate,
pH 4.4, v/v. Adjustment of the 1 m aqueous NH4formate to
pH 4.4 was performed with formic acid prior to adding the acetonitrile. The
eluate was split 90:10 between a fraction collector (90%) and an evaporative
light scattering detector (10%) (PL-ELS-1000, Polymer Laboratories). The
eluate was also monitored by measuring UV absorbance at 215 nm. Chromatography
was performed on an AKTA system (Amersham Biosciences) collecting 0.3-ml
fractions. Appropriate OPS fractions were combined, dialyzed, concentrated by
rotary vacuum evaporation, and subjected to structural analysis. De-O-acylation of OPS and Conversion of N-Acetimidoyl to N-Acetyl
Groups—Portions (6-8 mg) of the OPS fractions purified by HPLC were
de- O-acylated by treating with 12.5% v/v aqueous ammonia for 12 h at
35 °C ( 27). This treatment
also resulted in quantitative conversion of the N-acetimidoyl groups
to N-acetamido groups
( 28,
29). The progress of the
reaction was monitored by 1H NMR and MALDI-TOF MS described below.
The reactions were dialyzed versus water, then concentrated by rotary
vacuum evaporation, and subjected to analysis. Preparation and Fractionation of Oligosaccharide
Subunits—OPS samples were subjected to graded acid hydrolysis to
establish conditions giving a maximal yield of fairly large oligosaccharides,
with minimal yield of monosaccharides and minimal amounts of unhydrolyzed
polymer. Typically, for the de-O-acylated polysaccharide, 0.1
m trifluoroacetic acid at 105 °C for 1.5 h was most effective,
whereas for the native polymer, which was extensively O-acetylated, 3
h was required to obtain maximum yield of oligosaccharides. The products were
evaluated by size exclusion chromatography (SEC) on a Superdex-Peptide HR
10/30 FPLC column (Amersham Biosciences) eluted with 50 mm ammonium
acetate, pH 6.0. The oligo- and monosaccharide products were monitored by
evaporative light scattering detection with a 9:1 split. Semi-preparative
amounts of oligosaccharides were generated by hydrolyzing 2-4 mg of
polysaccharide per chromatographic run. Fractions were collected (0.4 ml), and
the saccharides were subjected to structural analysis. Glycosyl Analyses—Carbohydrate compositions of the OPS and
derived fractions were determined by preparing the TMS methylglycosides with
GC-MS (electron impact) analysis
( 13,
30) using a 30-m DB-5 fused
silica capillary column (J & W Scientific). Carbohydrate identities and
the locations of endogenous O-methyl groups were also determined by
GC-MS analysis of the alditol acetates, using a 30-m SP-2330 capillary column
(Supelco). Glycosyl residues containing carboxyl groups were analyzed by
preparing carboxyl-reduced alditol acetates by converting the carboxyl groups
to methyl esters (1 m methanolic HCl, 80 °C for 2 h), followed
by carboxyl reduction with NaBD 4 in water, 2 m
trifluoroacetic acid hydrolysis, and conversion to the alditol acetates
( 13,
31). Authentic
N-acetylquinovosamine (QuiNAc) was obtained from the R. etli
CE3 OPS ( 30). Authentic
6-deoxytalose ( l-isomer) was obtained from a streptococcal cell
wall glycan ( 32). Where
possible, the absolute configuration of glycosyl residues was determined by
preparing the diastereomeric TMS (-)-2-butylglycoside derivatives
( 33). GC-MS analysis was
performed on a DB-1 column with comparisons to authentic d-Fuc and
l-Fuc. Linkage analysis of neutral sugars was performed by permethylation
(Hakomori method), conversion to the PMAAs
( 13,
30), and GC-MS analysis. The
3-deoxy- d- manno-oct-2-ulosonic acid (Kdo) and other acidic
residue linkages were identified by sequential permethylation, reduction of
the carboxymethyl groups with lithium triethylborodeuteride (Aldrich), mild
hydrolysis (0.1 m trifluoroacetic acid, 100 °C, 30 min) to
cleave ketosidic linkages, reduction (NaBD 4), and conversion to the
PMAAs and GC-MS analysis ( 34).
Methylations using trideuteriomethyl iodide were also performed to confirm
sugar identities and location of endogenous O-methyl ether groups.
Oligosaccharide subunits derived from the OPS were analyzed as the
permethylated oligosaccharide alditols by reduction of the reducing end with
NaBD 4, followed by methyl esterification/permethylation (Hakomori
method). The products were analyzed by electron impact GC-MS (using SP-23230
and DB-5 columns), and by chemical ionization (CI) MS, using a 30-m DB-1
column with ammonia as reactant gas. Mass Spectrometry—Samples of OPS and derived
oligosaccharides were analyzed by matrix-assisted laser desorption ionization
(MALDI) mass spectrometry, using a Voyager-DE time of flight (TOF)
spectrometer (Applied Biosystems, Boston) in the positive and negative modes,
using a matrix of 100 mm 2,5-dihydroxybenzoic acid in 90% methanol.
The instrument was operated at an accelerating voltage of 25 kV with
extraction delay time of 200 ns. Samples were desorbed with a nitrogen laser
(λ = 337 nm) and the detector sensitivity was 1000 mV full scale. Mass
spectra were recorded over a m/z range of 500-20,000; spectra are the
summation of 200 acquisitions. Maltooligosaccharides (degree of polymerization
3-15) were used for calibration. Electrospray ionization-Q-TOF (ESI-Q-TOF) MS
analysis was performed on a Q-TOFII instrument (Micromass, Manchester, UK)
equipped with an electrospray source. Samples were infused into the nebulizer
at 5 μl/min, using nitrogen as the nebulization gas, and spectra were
collected in the positive ion mode. The predicted molecular mass of the
various saccharides was calculated using the following average incremental
mass values, based on the atomic weights of the elements: hexose, 162.142;
Kdo, 220.179; anhydro-Kdo, 202.164; 6-deoxyhexose, 146.143;
mono-O-methyl-6-deoxyhexose, 160.170;
di-O-methyl-6-deoxyhexose, 174.197;
2-N-acetamido-2,6-dideoxyhexose, 187.196;
3-N-acetamido-3-deoxyhexuronic acid, 217.178; acetimidoyl 41.052;
free reducing end, 18.015. Nuclear Magnetic Resonance Analyses— 1H spectra
and two-dimensional homo- and heteronuclear spectra of the OPS and derived
oligosaccharides were recorded at 25 °C on a Varian Inova 600- or 800-MHz
spectrometer, using a 5-mm triple probe and the standard Varian software
(Varian Medical Systems, Palo Alto, CA). Polymeric samples were analyzed on
the 800-MHz instrument, whereas derived oligosaccharides were analyzed on the
600-MHz instrument. Polysaccharides were dissolved in D 2O yielding
clear solutions at ~5 mg/ml; spectra were referenced to internal
2,2-dimethyl-2-silapentane-5-sulfonate sodium salt (δ H 0.00
ppm). Oligosaccharides were analyzed at 300-500 μg/260 μl of
D 2O using 5-mm symmetrical microtubes matched for D 2O
(Shigemi Inc., Allison Park, PA). In most experiments pre-saturation was
applied to the residual HDO signal. 1H- 1H COSY
( 35) data were recorded in the
absolute value mode with a 3.7-kHz spectral width and a matrix size of 512
× 4096 complex data points with eight scans per increment or with a 1024
× 4096 matrix and 16 scans/increment. 1H- 1H TOCSY
( 36) was recorded with a
mixing time of 80 ms and two sets of 256 time increments at 16 scans per
increment. Carbon-proton one-bond correlations were collected in the
1H detection mode with a gradient-selected
1H- 13C HSQC
( 37) with an acquisition time
of 0.2 s, collecting two arrays of 256 increments at 88-144 scans/increment.
The carbon spectral width was 16.5 kHz. Phase-sensitive
1H- 13C HMBC spectra were acquired with 256 × 2048
complex points at 96 to 144 scans/increment. The acquisition time was 0.27 s
( t2). Phasesensitive 1H- 1H ROESY
( 38) was collected with a
200-ms mixing time and matrix size identical to that of COSY and TOCSY, with
64 scans per increment. The anomeric configurations of the glycosyl linkages
were assigned from carbon-proton coupling constants
( JC1,H1) measured for the native OPS and derived
oligosaccharides by 1H- 13C HSQC analysis without
13C decoupling. Proton-proton coupling constants
( JH,H) were determined where possible from 1H
spectra, and by excitation of selected protons in a series of onedimensional
TOCSY experiments with mixing times from 0.08 to 0.12 ms. Isolation of LPS—The majority of LPS obtained by
phenol/water extraction of cultured R. leguminosarum 3841 cells was
recovered in the water layer (yield ~257 mg of total LPS/20 g dry cell
weight). The LPS was purified as described and analyzed by gel electrophoresis
(supplemental Fig. S1). The most abundant component was identified as a smooth
LPS (containing O-polysaccharide), having an apparent molecular mass centered
around 6,000 Da, of slightly lower mass than that of R. etli LPS
( 30) (supplemental Fig.
S1). Fractionation, Molecular Mass, and Composition Analysis of
OPSs—The polysaccharides derived from water layer LPS were
recovered at the void volume of Bio-Gel P-10 as described and analyzed by
positive ion ESI-Q-TOF-MS (). Five major polysaccharide components were detected (components
A-E) and are indicated along with their respective charge states. Each
polysaccharide, with the exception of component E, consisted of a family of
(pseudo)molecular ions differing in mass by 14 and 42 mass units, consistent
with variable content of endogenous O-methylation and
O-acetylation. Components A and B appeared to represent two forms of
a single polysaccharide, differing by the presence of a single anhydro-Kdo
residue (202.2 mass units), and components C and D represented two forms of a
secondary polysaccharide, also differing by a single Kdo (likely located at
the reducing end of the polysaccharides in each case, indicating the site of
core attachment) ( 13,
30,
34). Components A and B were
found to differ from the components C and D pair by 712 mass units, and
because no combination of commonly encountered glycosyl residues was evident
to relate this mass difference, the presence of two structurally unrelated OPS
was considered. Subsequently, experiments described below revealed the
presence of a new glycosyl residue, ultimately resulting in a repeating unit
mass of 712, and indicating that components A and B and C and D are in fact
the same polysaccharide, each pair differing by a single repeating unit. The polysaccharide mixture (A-E) was fractionated by HPLC as described
(supplemental Fig. S1). Three peaks were obtained, identified by MALDI-TOF MS
(supplemental Fig. S2) as peak 1 (component E), peak 2 (components B and D),
and peak 3 (components A and C). Glycosyl analysis revealed that HPLC peak 1
(component E) was a neutral polysaccharide composed of xylose, mannose, and
glucose, unrelated to the other components (details in supplemental material).
HPLC peaks 2 and 3, which included over 90% of the total peak area, contained
3Me6dTal and QuiNAc in a 0.8:1 ratio, along with lesser, non-stoichiometric
amounts of (ratios): fucose (0.19), 6-deoxy-3,4di- O-methyltalose
(3,4Me6d-Tal, (range 0.15-0.20), and an unidentified carbohydrate component
(0.2), which showed anomalous behavior during all derivatization procedures.
The configuration of the 6-deoxy-3- O-methylhexose residue was
assigned on the basis of retention times, identical to derivatives prepared
from authentic 6-deoxytalose
( 30,
32) when derivatized by either
the TMS methylglycoside, alditol acetate, or partially methylated alditol
acetate procedures. Linkage analysis of the peak 2 and peak 3 polysaccharides
yielded terminal-3Me6dTal and 3-linked QuiNAc in a 1:1 ratio (not shown).
Minor amounts of several fucose PMAA derivatives were again detected in
amounts not exceeding 15% of the 3-QuiNAc derivative. When the
carboxyl-reduced PMAAs were prepared following standard procedures, only these
same derivatives were again observed, along with derivatives of 4-linked Kdo.
These results were anomalous, because such derivatives in the detected ratios
( e.g. the absence of branch point residues) could not yield a
polysaccharide. The continued presence of nonstoichiometric amounts of fucose,
even after extensive chromatographic purification, and the presence of
anomalous derivatives all indicated that an unidentified glycosyl component
(or components) was present. Depolymerization of the OPS and Fractionation of Derived
Oligosaccharides—NMR spectroscopy of the primary OPS (HPLC peaks 2
and 3) yielded complex spectra inconsistent with the presence of a
diheteroglycan ( i.e. one composed only of 3Me6dTal and QuiNAc as
suggested by GC-MS analyses). Attempts were therefore made to obtain
structural subunits, and the major OPS fractions ( e.g. HPLC peak 3,
polysaccharides A and C) were subjected to various treatments to effect
partial and specific cleavage. Previously it was found that treating R.
etli OPS with base ( e.g. 0.25 m NaOH, 38 °C, 18
h) yielded oligosaccharides of defined size (primarily tetra- and
hexasaccharides), because of β-elimination of uronosyl residues
( 30). Similar alkali
treatments were tried with the 3841 OPS; however, the products consisted
mainly of monosaccharides and unsaturated degradation products, too small for
structural use. The base lability of the 3841 OPS thus appeared to be even
greater than that of R. etli OPS, suggesting the presence of
carboxylated or otherwise base-labile residues. However, mild acid was found
effective in releasing oligosaccharides of sufficient size to yield useful
structural information. Products thus obtained were fractionated on a Superdex
FPLC column supplemental Fig. S3), yielding two major oligosaccharide
fractions aligning in size with maltoheptaose (G7) and maltotetraose (G4).
Because it was already known that the polysaccharide contained endogenously
O-methylated residues in addition to 6-deoxy and amino sugars
( e.g. N-acetylquinovosamine), it appeared likely that the
oligosaccharide products were of a lower degree of polymerization than 7 or 4,
because the presence of 6-deoxymethyl groups, N-acyl, and
O-methyl ether groups all caused a significant increase in the
mobility of sugars during SEC compared with the parent sugars
( 39). Structural Analysis of Derived Oligosaccharides and Identification of a
New 3-Amino-3-deoxyhexuronic Acid Residue—Superdex fractions
aligning with the G7 and G4 standards, and also a lower mass fraction
corresponding to monosaccharides, were isolated and analyzed. The smaller
saccharide (“G4”), yielded a MALDI spectrum having ions at
m/z 381.5, 404.5, and 426.6, suggesting identities of M +
H+,M + Na+, and MNa + Na+ adducts
(supplemental Fig. S4A). Subjection of G4 to standard methanolysis
conditions and analysis of the TMS methylglycosides produced only a small
amount of fucose and a much larger amount of an unidentified component with a
late retention time in the range of a disaccharide or higher carbon sugar.
Similarly, MALDI analysis of the higher molecular mass saccharide (migrating
near “G7”) indicated a component 187 mass units higher than the G4
saccharide, suggesting that the G7 compound consisted of G4 linked to QuiNAc
(incremental mass 187.2, supplemental Fig. S4B). Composition analysis
confirmed this, revealing QuiNAc, in addition to a smaller, non-stoichiometric
amount of fucose and a much larger amount of the same late moving unidentified
component detected in G4. Assuming that both oligosaccharides contained
fucose, it was calculated that G4 must consist of fucose plus an unidentified
component having an incremental mass of 217 Da, whereas G7 presumably
consisted of this same oligosaccharide linked to QuiNAc. Because a mass of 217
Da is rather low to be a higher carbon glycosyl residue, it was determined
that the late moving peak observed during GC-MS analysis was in fact an
oligosaccharide that resisted methanolysis, most likely because of the
presence of an unidentified residue having a mass of 217 Da. If this residue
was glycosidically linked to fucose, the proposed acid stability of the
linkage would also account for the curious low recovery of Fuc during
composition analysis. The 1H NMR spectrum of the G4 compound identified three anomeric
signals in the ratio 1.0:0.8:0.2, suggesting a disaccharide in which the
reducing end residue existed in an α/β mixture (supplemental Fig.
S5). One N-acetyl signal was assigned (δH 2.05)
indicating that the disaccharide probably contained an amino sugar even though
QuiNAc was not detected during composition analysis. Signals for C6 methyl
protons were subsequently assigned to fucose on the basis of weak scalar
coupling between H5 and H4 protons. A 1H-13C HSQC
analysis ( and
supplemental Fig. S6) confirmed three anomeric centers and a single
nitrogen-bearing carbon with distinct upfield shift at δC
56.76. The 1H-1H COSY
() and TOCSY analyses
together with their carbon assignments defined the three glycosyl systems
(supplemental Table 1). Fucose was assigned as the reducing end residue, in
α-(δH1 5.22, 20%) and β-forms
(δH1 4.59, 80%). A downfield shift in carbons at δ
81.40 and δ 80.37 was assigned from COSY to C4 of the α- and
β-fucose residues, respectively, indicating that Fuc was substituted at
O4 by the new amino sugar. This was confirmed by 1H-13C
HMBC analysis (), which
revealed inter-residue correlations between Fuc H4 and an anomeric carbon at
δC 103.69, and between Fuc C4 and the corresponding anomeric
proton of the new residue (δH1 4.59). The COSY and HSQC
analyses of the new aminoglycosyl residue indicated that carbon C3 possessed
the upfield shift, and thus the amino group
(). However, C2 had a
normal shift (δ 71.80), consistent with an oxymethine carbon bearing an
unsubstituted hydroxyl group. In addition, only five carbon-proton pairs were
identified in this ring system. Selective excitation of the δ 3.54
proton, assigned from COSY as H4, yielded the one-dimensional TOCSY spectrum
(supplemental Fig. S5) in which all five protons of this spin system are
evident. The 1H-13C HMBC analysis
() revealed additional
correlations between H4 of the new residue and a downfield carbon at
δC 175.74, and between H5 of the new residue and this same
downfield carbon. These connectivities indicated that the new residue
contained a carbonyl carbon at C6, in addition to a nitrogen-bearing carbon at
C3, suggesting a 3-aminohexuronic acid. The presence of a carboxyl group at C6
would result in a mass increase of 30 mass units over that of a primary methyl
group, consistent with the suggested incremental mass of 217 mass units for
the new residue based on the MALDI MS analysis. The G4 compound was reduced with borodeuteride at the “reducing
end,” followed by permethylation and methyl esterification. Analysis of
the products by chemical ionization GC-MS yielded the CI spectrum
(supplemental Fig. S7) consistent with a permethylated disaccharide having (M
+ NH4)+1 m/z 515, in which the permethylated
hexosaminuronic acid occupies the nonreducing end. A separate portion of the
disaccharide was reduced at the reducing end, and the products were subjected
to the standard methanolysis procedure with preparation of TMS
methylglycosides. GC-MS analysis revealed a small amount of fucitol, instead
of the previously observed fucose (not shown), confirming that fucose occupied
the reducing end. 1H-1H ROESY analysis of the disaccharide (not shown)
identified an inter-residue NOE between H1 of residue A
(hexosaminuronic acid), and H4 of both the α/β-anomers of residue
B (Fuc). The relative intensity of the NOEs
(AH1/BβH4 intense; AH1/BαH4 weak), was
consistent with the relative abundance of the α/β-fucose anomers
and confirmed the glycosidic linkage of the hexosaminuronic acid to O4 of Fuc.
Intraresidue NOEs were observed for the β-anomer of the fucosyl system
between H1/H3, H1/H5, and H3/H5, consistent with axial orientation of these
protons and a 1C4 chair conformation for the
β-fucopyranose residue (assuming an absolute configuration of
l-, see below). The large JH,H coupling
constants for H1, H2, and H3 (supplemental Table 1) are also consistent with a
trans-diaxial arrangement of these protons. For residue A, the
location of the N-acetyl group at C3 was substantiated by an NOE
between the acetyl group protons and H3. The new residue also showed intense
NOEs between H1/H3, H1/H5, H3/H5, and H2/H4, consistent with an axial
orientation of protons and the pyranosidic 4C1
chair conformation for the β-anomer, assuming an absolute configuration
of d-. The intensity of COSY interactions
() and the scalar
JH,H coupling constants (9-10 Hz, supplemental Table 1)
indicated that all protons in this system were trans-diaxial,
consistent with the gluco-configuration. The anomeric configuration
was assigned as β-, from the JC1,H1 coupling constant
(164.63 Hz, supplemental Table 1), consistent with the δC,
δH, and JH,H values. Based on these data,
the new amino sugar was identified as
3-acetamido-3-deoxy-d-gluco-hexuronic acid (Glc3NAcA). A second set of inter-residue NOEs was observed between AH1 and
BH6 of both the α- and β-anomers of residue B
(fucose), suggesting that a favored conformation for this disaccharide
involved extensive rotation of the glycosidic linkage, such that the fucosyl
residue is flipped with respect to Glc3NAcA. Examination of model disaccharide
libraries (e.g. on line at Glycosciences and at CNRS) shows that the
disaccharides β-d-3-acetamido-3-deoxyglucopyranose
(1→4)-α-l-fucopyranose, and
β-d-glucopyranuronic acid-(1→4)-l-fucose, both
of which closely approximate the new disaccharide, exhibit several low energy
conformers, the lowest having ϕ = ψ = 99.9°, bringing Fuc H6 into
close proximity with H1 of (GlcA). An analogous NOE, between H1 of Glc3NAcA
and H6 of Fuc, was consistently observed in the polymeric O-chain as described
below. The 1H spectrum of the larger oligosaccharide (migrating in the
G7 region during SEC) revealed four anomeric signals, indicating that it was
probably a trisaccharide in which the reducing end existed in α/β
equilibrium (supplemental Fig. S5). 1H-1H COSY, TOCSY,
1H-13C HSQC, HMBC, and NOE analyses defined the four
glycosyl systems (supplemental Table 1) and identified the trisaccharide as
α-l-QuipNAc-(1→4)-β-d-Glcp3NAcA-(1→4)-α/β-l-Fucp-(1→,
in which the disaccharide identified above is substituted by a QuiNAc residue.
The 1H-13C HMBC spectrum is shown in
, and supporting data are
described in supplemental Experimental Procedures and Fig. S6. A remaining anomaly was the continued low yield of fucose during both the
standard methanolysis and alditol acetate procedures, presumably because of
the acid-stable glycosidic linkage of the new amino sugar. With the
identification of a hexosaminuronic acid, typified by extreme acid resistance,
samples of the disaccharide and OPS were subjected to hydrolysis in 4 or 6
m HCl, and the products were analyzed both as alditol acetates and
by methanolysis with conversion to the TMS methylglycosides. Both procedures
resulted in high yields of the new monosaccharide, as well as improved
recovery of fucose, which, although relatively acid-labile, was never the less
obtained in higher yields as a result of essentially quantitative cleavage of
the linkage. The electron impact-MS of the alditol acetate derivative is shown
in supplemental Fig. S7, and the GC-MS of the TMS methylglycosides are shown
in supplemental Fig. S8. With conditions identified to obtain fucose in
reasonable yield, the diastereomeric (-)-2-butylglycosides were prepared from
the disaccharide and OPS. GC-MS analysis identified derivatives identical to
those from authentic l-fucose, indicating that fucose was the
l-isomer. Analysis of the De-O-acylated Polysaccharide—Mass
spectrometry (Figs. and
), and 1H
NMR spectra (supplemental Fig. S9) indicated that the native OPS was highly
O-acetylated. To collapse the signals, portions of the native OPS
(from HPLC peak 3) were de-Oacetylated and analyzed by MALDITOF MS.
Two major molecular species were detected
(), having
pseudomolecular ions at m/z 2352 and 3064, differing by 712 mass
units. Each molecular species included a family of ions, differing by
±14 mass units, attributed to O-methyl heterogeneity involving
one or more of the endogenously O-methylated glycosyl residues
(i.e. 3Me6dTal and 3,4Me6dTal). Other structural features, including
two N-acetyl groups (supplemental Table 1) were assigned from the
1H NMR spectrum (supplemental Fig. S9). Identification of Glycosyl Systems and
Linkages—1H-1H COSY, TOCSY, and
1H-13C HSQC analyses identified four major and two minor
glycosyl systems comprising the de-O-acylated OPS (supplemental Table
1). A suitable starting point was identification of the three glycosyl
residues comprising the component trisaccharide QuiNAc
(α1→4)Glc3NAcA(β1→4)Fuc. The unique chemical shifts of
the Glc3NAcA system were identified, followed by QuiNAc; nitrogenbearing
carbons at δC 58.96 and 56.00 were assigned to C3 and C2 of
Glc3NAcA and QuiNAc, respectively. The C6 carbonyl (δC 176.04) of
Glc3NAcA was identified from correlations between H4/C6 and H5/C6 during
1H-13C HMBC analysis. Subsequently, the fucosyl spin
system was identified and distinguished from that of the 3Me6dTal system,
which had similar δC, δH, and
JH,H values. Initially, the distinction was assisted by
the identification of inter-residue HMBC correlations between the anomeric H/C
pair of the Glc3NAcA residue, and the corresponding C4/H4 pair of the
glycosidically linked (aglycon) residue, previously identified as fucose from
the oligosaccharide analysis. Partial HMBC spectra are shown in
. The data obtained thus far were consistent with three possible glycosyl
sequences for the polysaccharide repeating unit, shown in Structures
and
. To distinguish these possibilities, inter-residue HMBC correlations were
identified between H1 of 3Me6dTal and C3 of Fuc and between C1 of 3Me6dTal and
H3 of Fuc, indicating that 3Me6dTal residues were linked to O-3 of Fuc
residues (). Another set
of inter-residue correlations between Fuc H1-QuiNAc C3 and Fuc C1-QuiNAc H3
identified the remaining linkage, indicating a tetrasaccharide repeating unit
with fucose as the branch point residue (in Structure 1). The anomeric
configurations of the Fuc and 3Me6dTal residues were assigned as α-,
based on the JC1,H1 values (both > 170 Hz), consistent
with δH and δC values (measured in the
native OPS;supplemental Table 1). The δC values for the
fucosyl carbons are consistent with a pyranosidic form for fucose in the
polymer. This was supported by a strong intra-residue NOE between Fuc H3/H5
(). Two minor glycosyl systems were identified (supplemental Table 1), one was
a Kdo residue, which exists as a 2,7-anhydrofuranose and occupies the reducing
end of the polysaccharide. A second was identified as a variant of the
Glc3NAcA spin system, penultimate to Kdo, glycosidically linked to O4 of the
anhydro-Kdof residue. The assignment of these residues from COSY,
HSQC, HMBC, and NOE data is described in the supplemental material. The
glycosidic sequence at the nonreducing end of the de-O-acyl OPS was
deduced from the MALDI spectrum (). Consecutive losses of 160 and 146 mass units are
consistent with loss of 3Me6dTal and Fuc, indicating that the nonreducing end
terminates in the sequence 3Me6dTal→ Fuc→. The major ion
(m/z 2352) is consistent with three repeats of the tetrasaccharide
repeating unit (in addition to one anhydro-Kdo residue) for the main
de-O-acylated OPS (corresponding to de-O-acylated component
A, identified in ). NOE Analysis and Conformational Features of the De-O-acyl
OPS—The 1H-1H ROESY spectrum of the
de-O-acylated OPS yielded inter-residue NOEs from all anomeric
protons (), confirming
the linkages and glycosidic sequence assigned from HMBC correlations. Several
longer range, inter-residue NOEs provided additional insight into several
aspects of polysaccharide conformation. An NOE (supplemental Fig. S10) between
A1/B6 was analogous to that observed in the oligosaccharides,
indicating that a favored conformation exists in which the fucosyl residue is
flipped over with respect to Glc3NAcA. Another NOE was observed between
B4/D5 (Fuc H4-3Me6dTal H5) requiring that the plane of the
3Me6dTal ring is rotated out of the plane of, and nearly perpendicular to, the
Fuc ring. Models show that only α-linked-3Me6dTal residues, and not
β-, could form such an NOE with Fuc, consistent with the anomeric
assignment from JC,H and δC values
(supplemental Table 1). Moreover, models show clearly that if the absolute
configuration of the α-3Me6dTal residue were l-, then
formation of a B4/D5 NOE would necessitate an additional NOE
between B3/D3; however, the latter was not observed. Only the
α-d-isomer of 3Me6dTal would allow the formation of the
B4/D5 NOE with-out the accompanying B3/D3 NOE,
indicating that the α-3Me6dTal residues have the d-absolute
configuration. Another interesting feature is the arrangement of the side
chain 3Me6dTal residues. An inter-residue NOE was observed between the
N-acetyl group methyl protons of residue C (QuiNAc) and H5 of
residue A (Glc3NAcA). The presence of this NOE, and an NOE between
C1/A4, would suggest that the two N-acyl groups (of
residues A and C) be on “opposite sides” of the
polymer, with residue A flipped with respect to residue C
(). This arrangement
would necessitate that the side chain 3Me6dTal residues be distributed on
“alternating sides” of the polysaccharide. Models also show that
only the l-configuration for QuiNAc would allow the simultaneous
formation of these two NOEs (assuming the d-configuration for
GLc3NAcA). Localization of O-Acetyl and N-Acetimidoyl Groups and NOE Analysis of
the Native OPS—The presence of N-acetimidoyl groups in the
native PS was evident from the characteristic δ C and
δ H values for this moiety ( C = O, 166.84;
C H3, 2.25/2.21, see supplemental Table 1). HMBC
correlations between the carbonyl carbon and H3 of residue A, and
between this carbonyl carbon and the methyl group protons (δ H
2.25/2.21) allowed assignment of the N-acetimidoyl group to C3 of
residue A (Glc3NAmA). Confirmation of the presence and location of this
moiety was obtained from the large upfield shift of the nitrogen-bearing
carbon (C3), from δ C 62.81 in the native PS to 58.96 in the
de- O-acylated PS (supplemental Table 1). The mild
de- O-acylation conditions typically cause quantitative conversion of
N-acetimidoyl to N-acetyl groups, and the resulting carbon
shift is typically-3 ppm or more
( 28,
29,
40). Comparison of the native spectra with those of the de- O-acylated
OPS revealed four O-acetyl groups, identified from the downfield
δ H shift of four ring protons into the anomeric region,
indicative of de-shielded protons attached to carbons bearing the
O-acyl groups. COSY and HSQC analyses (supplemental Fig. S11)
indicated that most of the shifted protons were attached to C2 of 3Me6dTal
residues, indicating that each repeating unit was O-acetylated at C2
of the 3Me6dTal residue in the fully acetylated molecular species. A downfield
shift (δ C 1-3 ppm) was observed for each C2 carbon of the
3Me6dTal residues, relative to those of the de- Oacylated OPS
(supplemental Table 1), reflecting the expected α-effect because of
acylation ( 41). Also
consistent with C2 as the site of O-acylation was a substantial
upfield shift of the anomeric carbon of 3Me6dTal in the native OPS compared
with that of the de- O-acylated OPS, indicative of the β-effect
because of O-acylation at C2
( 41). The assignment of
downfield shifted protons to H2 of 3Me6dTal residues was supported by NOEs
between H1/H2 and H2/H3 of this residue (residue D) in the native OPS
(supplemental Fig. S12). Heterogeneity in each glycosyl system was observed in
the native OPS, particularly for the terminal 3Me6dTal residues (supplemental
Table 1). The occurrence of three distinct sets of
2- O-acetyl-3Me6dTal δ H/δ C signals
probably originates from the three repeating units, each unit giving rise to a
slightly different magnetic environment for the side chain 3Me6dTal residues.
Bulky O-acetyl groups at C2 presumably contribute to this
heterogeneity. A minor set of C2-shifted protons was assigned to fucosyl
residues (supplemental Table 1), indicating that one or two of the fucosyl
residues in the linear portion of the polysaccharide can also be
2- O-acetylated; however, the random or specific locations of
O-acetylated fucose ( i.e. which repeating units) was not
assigned. Further insight into the degree of O-acetyl heterogeneity
in the native polysaccharide can be deduced from the MALDI mass spectrum
(), which
indicates a maximum of four major molecular species reflecting varying degrees
of O-acetylation. The glycosyl sequence of the native OPS was confirmed by inter-residue
dipolar correlations from all anomeric protons (supplemental Fig. S12) and by
inter-residue 1H-13C HMBC couplings. A notable
difference between the native and de-Oacylated polysaccharides was
the presence of an unusual “capping” residue at the nonreducing
end of the former, identified from COSY, TOCSY, HSQC, and HMBC analyses as a
4,6-dideoxy-4-formamido-hexopyranose of unidentified configuration (residue
E, supplemental Table 1 and ). This terminal residue and the residue to which it is
glycosidically linked are apparently cleaved from the polysaccharide during
de-Oacylation. Additional details identifying this residue are in
supplemental materials. The complete structures of the OPSs from the
free-living forms of R. leguminosarum 3841 and R. etli CE3
are compared in . The primary O-polysaccharide produced by free-living R.
leguminosarum 3841 is a branched tetraheteroglycan having the structure
shown in . The
polysaccharide has a number of rather unusual structural features for an
O-antigen, including a uniform size (three to four repeats with little
variation in the number of repeating units), low molecular mass (~3000
Da), extensive O-acetylation and O-methylation of side chain
residues (6-deoxy-3- O-methyltalose, 17% of which are also methylated
at O4), and a backbone containing uncommon and/or previously unreported amino
sugars, including QuiNAc and a new 3-amino-3-deoxyhexuronate residue,
3-acetimidoylamino-3-deoxy- d- gluco-hexuronic acid
(Glc3NAmA, designated rhizoaminuronic acid). All previously described
hexosaminuronic acids in bacterial polysaccharides contain an amino group at
C2. Although unusual compared with enterobacterial O-antigens, similar
structural features are found in the OPS synthesized by the closely related
R. etli CE3, which produces a low molecular mass (3330 Da), uniform
size O-chain also containing side chains of 6-deoxy-3- O-methyltalose
and a capping residue of 2,3,4-tri- O-methylfucose
( 30). Other rhizobial O-chains that have been characterized, such as that from
Sinorhizobium sp. NGR234, contain endogenously O-methylated
l-rhamnose ( 6), and
the O-chain from Mesorhizobium loti NZP2213 contains
O-acetylated-Tal and 6-deoxy-2- O-methylTal
( 16), an otherwise infrequent
residue. Endogenously O-methylated and O-acetylated
6-deoxyhexose residues may confer a degree of hydrophobicity on these glycans,
a property that could influence bacteroid surface chemistry and help
facilitate symbiotic infection. In addition to the primary OPS, the free living form of R.
leguminosarum 3841 also produces a secondary polysaccharide
( and supplemental Fig.
S1), composed of xylose, mannose, and glucose. This neutral polysaccharide is
eventually separated from the primary OPS by HPLC (supplemental Fig. S1). This
xylomannan shows limited affinity to polymyxin-agarose, and preliminary
results indicate that it is not attached to a “normal” rhizobial
lipid A but rather is associated with an acyl glycerol
moiety. 3
Interestingly, the relative abundance of these two polysaccharides is modified
by external physiological
conditions. 4 In free
living cells cultured under normal conditions, the xylomannan constitutes only
5% (supplemental Fig. S1) of the total PS isolated from the LPS fractions, the
other 95% is the primary OPS. When cells are grown at pH 5 (conditions
intended to mimic those within the symbiosome), the xylomannan becomes the
dominant PS, constituting >95% of the total polysaccharide components. In
an earlier report ( 18),
composition analysis of LPS-associated carbohydrates extracted from cultured
R. leguminosarum 3841 cells showed an increase in xylose and mannose
when grown under the physiological extremes; however, it was not known if this
represented the appearance of a new polysaccharide or a structural alteration
to a single PS. This study shows that the source of xylose, man-nose, and
glucose is indeed a separate polysaccharide, and preliminary results indicate
that a similar or identical xylomannan is also synthesized by R.
leguminosarum 3841 bacteroids isolated from P. sativum nodules.
The structure of this putative “bacteroid-specific” xylomannan,
its regulation, and relation to lipid A-core is currently under study. Recent studies have succeeded in isolating bacteroids in sufficient
quantity and purity to allow direct examination of bacteroid-specific LPS,
providing new insight into the nature of the bacteroid surface and the role of
LPS in bacteroid survival and adaptation. In the Sinorhizobium sp.
NGR234, Vigna unguiculata model system, the transition from free
living cells to bacteroids is accompanied by a shift in LPS surface chemistry,
from a rough LPS lacking O-antigen in the vegetative state to a smooth LPS
composed of a rhamnan O-antigen attached to a structurally modified core-lipid
A ( 6). The endogenously
methylated rhamnan homopolymer is relatively hydrophobic, and the modified
core region lacks the acidic sugars commonly found in the antigenic outer core
of LPS from free living sinorhizobia. This “phase shift” in LPS
surface chemistry may promote proper interaction between the bacteroid and the
surrounding symbiosome membrane or attenuate the host innate immune response
in some way, possibly as a mimic of host structures
( 3- 6).
Interestingly, the NGR234 rhamnan O-antigen has the same primary sequence as
the A-band O-antigen of Pseudomonas aeruginosa, which is selectively
expressed during chronic cystic fibrosis lung infection, where it is
associated with an increased duration of infection
( 42). In the other well
studied system, that of R. etli CE3, bacteroid-specific LPS isolated
from R. etli CE3 bacteroids (from Phaseolus nodules) were
found to be structurally similar but not identical to the free-living R.
etli LPS ( 7). Here a
significant difference was an additional O-methyl group at the
2-position of one of the fucosyl residues in the O-antigen of the bacteroid
LPS. In R. leguminosarum 3841, the structures of the bacteroid LPS
and other surface components are not yet known. However, it was previously
found that RL3841 bacteroids and free-living (vegetative) cells could
be partitioned between dodecane and water, providing a relative estimate of
cell surface hydrophobicity
( 18). Vegetative bacteria
grown under conditions of low pH and oxygen, and bacteroids isolated from pea
nodules both adhered preferentially to the dodecane layer, whereas vegetative
cells grown under normal conditions favored the water layer, suggesting that
the entire bacteroid surface (and of cells grown under physiological extremes)
was more hydrophobic than that of normally cultured free-living cells. The majority of rhizobial LPS-structure mutants appear to be defective at
the later stages of infection, i.e. during and after the stage of
bacterial invasion into the nodule cells
( 2,
3,
16). Consequently, LPS are
likely to play an important role in later stage events such as endocytosis,
bacteroid survival, and interaction with plant symbiosome membrane components.
Two LPS-structure mutants, an lpsB mutant of Sinorhizobium
meliloti Rm1021 ( 43) and
the R. etli mutant CE166
( 13,
14), are both compromised in
their ability to induce normal nodules; however, the early stage events
( e.g. bacterial adhesion, colonization of root hairs, and infection
thread initiation) appear relatively unaffected. The lpsB mutant is
also sensitive to cationic antimicrobial peptides, components of the plant
innate immune system ( 43,
44). Parallels exist among
animal pathogens, and LPS mutants of Salmonella
( 45), Pseudomonas
( 46), and the intracellular
pathogen Brucella
( 47) all show increased
sensitivity to cationic antimicrobial peptides ( e.g. plant defensins)
resulting in reduced virulence and loss of intracellular survival
( 48). Related studies suggest
that LPS could act by several mechanisms, including passive protection, and
through active mechanisms, whereby LPS serve as signals specifically
recognized by plant receptors, allowing the infection to proceed
( 2- 5). The complete genome of strain RL3841 was recently sequenced
( 25), and preliminary
analyses 5 indicate
there appear to be two major gene clusters devoted to O-chain/LPS
biosynthesis, one of which is chromosomal and the other located on plasmid
pRL9, distinct from the symbiosis plasmid pRL10. The particular features of
this polysaccharide, specifically uniform size and low molecular mass, appear
to be most compatible with the “monomeric” biosynthetic mechanism,
in which individual monosaccharide residues are transferred consecutively from
the glycosyl donor (XDP-sugar) to the nonreducing end of the growing chain
(reviewed in Ref. 49). This
mechanism has been associated with the uniformly sized OPSs synthesized by
certain strains of Escherichia coli, Rhodospirillum rubrum, and
others ( 49). This type of
discrete size OPS is frequently encountered in bacteria possessing an ordered
surface typified by a crystalline glycoprotein surface layer ( i.e.
S-layer), including diverse Gram-negative eubacteria
( 50). R. leguminosarum/R.
etli are not known to possess an S-layer; however, the presence of OPS of
uniform size could allow the ordered assembly of other surface components or
promote interaction with plant-derived symbiosome membranes, either through
multiple weak ( e.g. hydrophobic) interactions, or via specific plant
receptors such as lectins ( 51,
52). Unlike rhizobial capsular polysaccharides, which often have a high negative
charge density ( 21,
53), surveys of O-chain
structures from diverse species seem to suggest that highly negatively charged
OPS are not a particularly favored structural feature
( 54,
55). A maximum charge density
of one negative residue per every three repeating unit residues is common, and
the incorporation of additional negative residues is frequently compensated
( 55) by the introduction of a
positive charged group, e.g. ethanolamine. The rhizobial O-chain
biosynthetic machinery seems to be particularly stringent in this regard.
Among published rhizobial structures, neutral OPS that have some degree of
hydrophobicity appear to be favored, and residues imparting net negative
charge are either absent ( 6,
16) or when present are
blocked by esterification (as in R. etli CE3) or neutralized with a
positive substituent ( N-acetimidoyl group, shown here for R.
leguminosarum) to yield the zwitterion. At normal physiological pH, at or
around neutrality, it would be expected that the RL3841 OPS has no
net charge, and this is the form believed to be expressed while the bacteria
adhere and colonize the plant surface. Following internalization, the
bacteroid surface/O-antigen could encounter acidic pH, such as the symbiosome
compartment ( 2,
22). If the pH approaches the
p Ka value of the uronic acid carboxyl ( e.g. pH
4.0), a net positive charge could exist on these OPS and at the bacteroid
surface; the OPS could behave transiently as a polycationic species, forming
an electrostatic barrier that would repel cationic antimicrobial peptides
( 4,
44). Hexosaminuronic acids occur rarely, and the absence of an amino group at C2
is a structural feature that raises biosynthetic questions. Several 2-amino
( 56,
57) and 2,3-diaminohexuronic
acids have been characterized, notably from the O-antigens of Vibrio
ordalii ( 58),
Bordetella hinzii
( 59), and several
Pseudomonas serotypes
( 55). In P.
aeruginosa, UDPGlcNAc was the common intermediate in the biosynthesis of
the 2,3-di- N-acylated hexosaminuronic acids, including
d-ManNAc3NAcA and d-ManNAc3AmA
( 60). Their biosynthesis
generally involves epimerases, followed by C6 dehydrogenation of the
l/ d-UDP-hexNAc to yield the UDP-hexosaminuronate. In the
case of 3-amino hexoses ( e.g. kanosamine) and
3-amino-3,6-dideoxyhexoses ( e.g. mycosamine), evidence indicates that
the nitrogen donor is glutamine, with an NAD-dependent transaminase acting on
the XDP-3-keto-sugar intermediate
( 61,
62). Thus, a possible route to
rhizoaminuronic acid could involve C6 oxidation of an XDP-3-aminohexose
intermediate. This unusual 3-amino sugar is a suitable candidate for selective
inhibition of OPS synthesis in R. leguminosarum 3841. We are grateful to John Glushka for advice with NMR analyses, Elmar
Kannenberg for helpful discussion, and Christopher Hackney for technical
assistance. The Complex Carbohydrate Research Center was supported in part by
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