From the Department of Biochemistry, Duke University
Medical Center, Durham, North Carolina 27710 and the
§ Department of Pharmacology and Molecular Sciences, The
Johns Hopkins University School of Medicine,
Baltimore, Maryland 21205
Received for publication, January 13, 2003
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ABSTRACT |
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The structures of Rhizobium
leguminosarum and Rhizobium etli lipid A are distinct
from those found in other Gram-negative bacteria. Whereas the more
typical Escherichia coli lipid A is a hexa-acylated
disaccharide of glucosamine that is phosphorylated at positions 1 and
4', R. etli and R. leguminosarum lipid A
consists of a mixture of structurally related species (designated A-E) that lack phosphate. A conserved distal unit, comprised of a diacylated glucosamine moiety with galacturonic acid residue at position 4' and a
secondary 27-hydroxyoctacosanoyl (27-OH-C28) as part of a 2'
acyloxyacyl moiety, is present in all five components. The proximal end
is heterogeneous, differing in the number and lengths of acyl chains
and in the identity of the sugar itself. A proximal glucosamine unit is
present in B and C, but an unusual 2-amino-2-deoxy-gluconate
moiety is found in D-1 and E. We now demonstrate that membranes of
R. leguminosarum and R. etli can convert B to
D-1 in a reaction that requires added detergent and is inhibited by
EDTA. Membranes of Sinorhizobium meliloti and E. coli lack this activity. Mass spectrometry demonstrates that B is
oxidized in vitro to a substance that is 16 atomic mass
units larger, consistent with the formation of D-1. The
oxidation of the lipid A proximal unit is also demonstrated by
matrix-assisted laser desorption ionization time-of-flight mass
spectrometry in the positive and negative modes using the model
substrate, 1-dephospho-lipid IVA. With this material, an
additional intermediate (or by product) is detected that is tentatively
identified as a lactone derivative of 1-dephospho-lipid
IVA. The enzyme, presumed to be an oxidase, is located
exclusively in the outer membrane of R. leguminosarum as
judged by sucrose gradient analysis. To our knowledge, an oxidase associated with the outer membranes of Gram-negative bacteria has not
been reported previously.
The Rhizobiacea are agriculturally important Gram-negative
bacteria that are able to establish a symbiotic relationship with the
root cells of certain plants (1). The symbiotic bacteria provide the
plant with NH4+ by fixing N2,
whereas the plant provides the bacteria with reduced carbon sources
(2). A detailed understanding of the many factors contributing to the
complex interplay between the plant host and the microbes is beginning
to emerge (3). A few of the key components needed for effective
symbiosis and nitrogen fixation include flavonoids (4), Nod factors (5,
6), receptor kinases (7), various exopolysaccharides (8, 9), cyclic
LPSs are macromolecular glycolipids present on the outer surfaces of
the outer membranes of Gram-negative bacteria (15-18). The structure
of LPS may be subdivided into the lipid A region, which embeds the LPS
in the outer membrane, the nonrepeating core oligosaccharide, and the
distal O-antigen polysaccharide. The effects of changing LPS structure
on symbiosis and nitrogen fixation are not fully characterized (14).
Mutants of Rhizobium leguminosarum and Rhizobium
etli that lack O-antigen are defective in the infection process,
producing poorly differentiated, nonfunctional nodules (13, 19-21).
Whether or not mutations in lipid A biosynthesis are deleterious to
symbiosis remains to be determined.
Recent studies of the LPS of R. leguminosarum and R. etli, which are distinct bacterial species based on their
ribosomal RNA sequences (22), have revealed that both bacteria possess
unusual lipid A molecules (23-25) and core oligosaccharides (14, 20, 26, 27). Although the classical lipid A structure, typified by that of
Escherichia coli, consists of a hexa-acylated disaccharide of glucosamine that is phosphorylated at positions 1 and 4' (15-18), R. etli and R. leguminosarum lipid A is recovered
as a mixture of structurally related components (A to E) that are not
phosphorylated (Fig. 1) (24, 25, 28). A
conserved distal unit, comprised of a diacylated glucosamine residue
with a secondary 27-OH-C28 acyl chain as part of an acyloxyacyl moiety
at position 2' and a galacturonic acid residue at position 4', is
present in all of the components (Fig. 1) (24, 25, 28). The
microheterogeneity of R. etli lipid A arises mainly in the
proximal unit, which may contain acyl chains of different lengths at
position 2 or may be deacylated at position 3 (Fig. 1). Although
glucosamine constitutes the proximal units of components B and C, its
oxidized form, 2-amino-2-deoxy-gluconate (2-aminogluconate), is
present in D-1 and E (Fig. 1) (24, 25, 28). Components C and E are
3-O-deacylated derivatives of the more abundant species B
and D-1, respectively (24, 25, 28). Component A (not shown in Fig. 1)
appears to be an elimination product generated from D-1 during the mild
acid hydrolysis procedure used to release lipid A from the LPS core
(24, 25). D-2 is an isomer formed from D-1 in a nonenzymatic reaction
and is attributed to acyl chain migration on the proximal unit (24,
25).
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-glucans (10, 11), K-antigens (12), and lipopolysaccharides
(LPSs)1 (13, 14).
View larger version (14K):
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Fig. 1.
Structures of the predominant lipid A species
in E. coli, R. etli, and R. leguminosarum. R. etli lipid A, which is
thought to be the same as that of R. leguminosarum, is a
mixture of several related species (23-25). In contrast to E. coli lipid A, all R. etli lipid A species lack
phosphate groups (24, 25). Instead, each one contains a galacturonic
acid moiety at position 4' and a single acyloxyacyl unit, featuring an
unusual 27-OH-C28 secondary acyl chain, at position 2' (24, 25). The
dashed bonds indicate partial substitutions in the major
R. etli components B and D-1 (24, 25). The molecular weight
of the largest, fully substituted form of each component is indicated
in parentheses. The minor R. etli
lipid A species C and E (not shown) are the 3-O-deacylated
derivatives of B and D-1, respectively (24, 25). The proximal sugar is
a glucosamine unit in B and an aminogluconate moiety in D-1 (24, 25).
The proximal
3-deoxy-D-manno-2-octulosonic acid (Kdo)
residue of the core oligosaccharide (not present in lipid A prepared by
mild acid hydrolysis) is attached at position 6' in intact LPS.
To date, the occurrence of the 2-aminogluconate moiety is limited to
the lipid A of R. etli and R. leguminosarum (23,
24). A plausible pathway for 2-aminogluconate formation could involve the oxidation of the proximal glucosamine unit of component B to
generate D-1. In conjunction with our recent re-evaluation of the
structures of R. etli and R. leguminosarum lipid A (25), we discovered that crude
extracts of R. etli and R. leguminosarum can
convert 14C-labeled component B to a lipid that migrates
with D-1 during thin layer chromatography. We now report the detailed
characterization of this novel oxidative reaction using both the
natural product B and the relatively simple model substrate,
1-dephospho-lipid IVA. Remarkably, the enzyme is an outer
membrane protein and is presumed to be an oxidase. These findings are
consistent with the idea that 2-aminogluconate formation is a late step
in R. leguminosarum lipid A biosynthesis. As demonstrated in
the accompanying manuscript, a novel gene, designated lpxQ,
encodes the putative oxidase.
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EXPERIMENTAL PROCEDURES |
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Materials-- Glass-backed 0.25-mm Silica Gel 60 thin layer chromatography plates were from Merck. Chloroform, ammonium acetate, and sodium acetate were obtained from EM Science. Pyridine, methanol, and formic acid were from Mallinckrodt. [U-14C]acetate was purchased from Amersham Biosciences.
Bacterial Growth Conditions and Membrane Preparations-- R. leguminosarum 3855 was grown at 30 °C in TY broth (5 g of tryptone and 3 g of yeast extract/liter) supplemented with 10 mM CaCl2. R. etli CE3 and Sinorhizobium meliloti 1021 were grown in TY broth supplemented with 10 mM CaCl2, 20 µg/ml nalidixic acid, and 200 µg/ml streptomycin sulfate. E. coli strains were grown at 37 °C in LB broth (29) with one of the following antibiotics, depending on the resistance markers of the plasmid that the strain harbors: ampicillin (100 µg/ml), tetracycline (15 µg/ml), and kanamycin (25 µg/ml). Table I describes the various bacterial strains used.
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Rhizobium cultures (0.5 to 1 liter) were grown to late log
phase (OD550 = ~1.0) and then harvested by
centrifugation at 6,000 × g for 15 min at 4 °C. All
subsequent steps were carried out at 4 °C. The cell pellets were
washed with 50 mM HEPES, pH 7.5, with a volume that
was 1/10th that of the original culture volume. The cells
were collected by centrifugation and resuspended in a minimal volume
(usually 5 ml) of 50 mM HEPES, pH 7.5, and stored at
80 °C if they were not immediately lysed. The frozen cells were
thawed, resuspended in 50 mM HEPES, pH 7.5, at a protein
concentration of ~3-10 mg/ml, and broken by three passages through
an ice-cold French pressure cell (SLM Instruments, Urbana, IL) at
18,000 p.s.i. Large cell debris and unbroken cells were removed by
centrifugation at 12,100 × g for 10 min. The membranes
were recovered from the cytosol by ultracentrifugation at 149,000 × g for 1 h. The pellet containing the membranes was
washed by homogenization in the same volume of 50 mM HEPES,
pH 7.5. After a second ultracentrifugation, the washed membranes were
homogenized in 50 mM HEPES, pH 7.5, at a protein
concentration of ~5-15 mg/ml. The samples were stored in aliquots at
80 °C. The protein concentrations were determined by the
bicinchoninic acid assay with bovine serum albumin as the standard
(30).
Subcellular Localization of the Lipid A Oxidase--
Subcellular
localization of the lipid A oxidase in R. leguminosarum 3855 membranes was determined using a protocol similar to that described by
Trent et al. (31). Briefly, a 1-liter culture of R. leguminosarum 3855 was grown with shaking (225 rpm) at 30 °C
for 16 h (OD550 = ~1.0). The culture was centrifuged
at 6,000 × g for 10 min at 4 °C. The cell pellet
was resuspended in 7.8 ml of 50 mM HEPES, pH 7.5, containing 0.5 mM EDTA, and the cells were lysed by passage
through a French pressure cell at 10,000 p.s.i. Crude cell-free extract
was prepared by removal of unbroken cells and large debris by
centrifugation at 12,100 × g for 10 min at 4 °C,
and the supernatant was recovered. One-half of the crude extract was
stored at 80 °C, whereas the remaining half was used to prepare
washed membranes, as described above. The washed membranes were
homogenized with a 25-gauge 1/2 syringe needle in a total volume of 2.5 ml of 50 mM HEPES, pH 7.5, containing 0.5 mM
EDTA. After being layered on top of a seven-step isopycnic sucrose gradient as described by Guy-Caffey et al. (32), the membranes were centrifuged in a swinging bucket rotor at 155,000 × g for 18 h at 4 °C. The fractions (0.5. ml) were
collected, and their protein content was determined by the
bicinchoninic acid assay, as described above. The following undiluted
fraction volumes were used for the various assays: 3 µl for the lipid
A oxidase reaction, which was terminated after 30 min; 50 µl for the
NADH oxidase assay (33, 34); and 20 µl for the phospholipase A assay
(35). The turbidity (OD600) of each fraction was measured to confirm the presence of membrane fragments. The activity for each
fraction was calculated as a percentage of the total activity throughout the entire gradient.
Preparation of Nonradioactive and 14C-Labeled R. etli Lipid A Component B-- Nonradioactive B was prepared from R. etli CE3 as described previously (24, 25). To make 14C-labeled component B, a 50-ml culture of R. etli CE3 was grown to A550 = 1 in the presence of 500 µCi of [U-14C]-acetate (50 mCi/mmol) (24, 25). A combination of DEAE-cellulose column chromatography and preparative thin layer chromatography (24) was used to purify [14C]B.
Preparation of 1-Dephospho-lipid IVA--
Lipid
IVA was isolated as described by Raetz et al.
(36) and further purified by reversed phase column chromatography (37). To cleave the phosphate at the anomeric position (38, 39), 3.2 mg of
lipid IVA in a 16 × 125-mm glass tube with a
Teflon-lined cap was resuspended by sonic irradiation in 3.6 ml of 0.1 M HCl and placed in a boiling water bath for 15 min. After
cooling, the hydrolyzed material was converted into a two-phase
Bligh-Dyer system (40) by the addition of 4 ml of CHCl3 and
4 ml of MeOH. After mixing, the solution was centrifuged in a clinical
centrifuge for 10 min. The lower phase was removed and transferred to a
clean glass tube. The upper phase was re-extracted once with 4 ml of a
fresh lower phase of a pre-equilibrated two-phase Bligh-Dyer system
(40). The lower phases from both extractions were pooled and dried down
under a stream of N2. The dried product was then redissolved in 450 µl of CHCl3/MeOH (4:1, v/v) and
spotted in a line 20 mm from the edge of a 20 × 20-cm silica TLC
plate. The plate was allowed to dry and developed in CHCl3,
pyridine, 88% formic acid, H2O (50:50:16:5, v/v) until the
solvent front was ~5 cm from the top of the plate. The solvents were
removed by drying the plate with a cold air stream for 30 min. The band
of interest, which is transiently visible as a white zone during the
drying process, was localized by placing the plate on top of a light
box. The band was outlined in pencil and scraped off with a clean razor
blade. The silica chips were transferred into a 16 × 125-mm glass
tube equipped with a Teflon-lined cap. The lipid was extracted from the
chips by resuspending them in 3.8 ml of an acidic single-phase
Bligh-Dyer solution, consisting of CHCl3, MeOH, 0.1 M HCl (1:2:0.8, v/v), followed by brief sonic irradiation.
The solution was then converted into a two-phase Bligh-Dyer mixture by
adding 3 ml of CHCl3, 2 ml of MeOH, and 2.8 ml of distilled
H2O. After thorough mixing and centrifugation for 8 min in
a clinical centrifuge, the lower phase was recovered and passed through
a Pasteur pipette plugged with glass wool. The extraction of the chips
was repeated once, and three drops of pyridine were added to the pooled
lower phases to neutralize any residual HCl. After drying the lower
phases with a stream of N2, the sample was stored at
20 °C.
Further purification of the 1-dephospho-lipid IVA was done by passing the material from the preparative TLC step through a 0.5-ml DEAE-cellulose column (Whatman DE-52), previously equilibrated as the acetate form in CHCl3, MeOH, H2O (2:3:1, v/v) (24, 36, 41). The above sample was redissolved in 12 ml of CHCl3, MeOH, H2O (2:3:1, v/v) and loaded onto the column. The flow-through was collected as one fraction. The column was washed with 1 ml of CHCl3, MeOH, H2O (2:3:1, v/v), and stepwise elution was achieved by including increasing ammonium acetate concentrations in the solvent mixture (24, 36). The column was thus successively washed with 2 ml of CHCl3, MeOH, 30 mM aqueous NH4Ac (2:3:1, v/v), 2 ml of CHCl3, MeOH, 60 mM aqueous NH4Ac (2:3:1, v/v), 4 ml of CHCl3, MeOH, 120 mM aqueous NH4Ac (2:3:1, v/v), 2 ml of CHCl3, MeOH, 240 mM aqueous NH4Ac (2:3:1, v/v), and finally with 2 ml of CHCl3, MeOH, 500 mM aqueous NH4Ac (2:3:1, v/v). 0.5-ml fractions were collected throughout. Elution of the lipid from the column was monitored by TLC using silica gel plates (5 × 10 cm), which were developed with the solvent CHCl3, pyridine, 88% formic acid, H2O (50:50:16:5, v/v). Some of the 1-dephospho-lipid IVA emerged from the column in the late CHCl3, MeOH, 60 mM NH4Ac wash, but most of the material eluted in the CHCl3, MeOH, 120 mM NH4Ac step. Only the fractions from the 120 mM wash were processed further as the source of 1-dephospho-lipid IVA substrate for in vitro incubations with the R. leguminosarum membranes. The selected fractions were pooled and converted into a two-phase Bligh-Dyer system, consisting of CHCl3, MeOH, H2O (2:2:1.8, v/v). The solvents were mixed and centrifuged for 8 min at room temperature in a clinical centrifuge. The upper phase was re-extracted once with a pre-equilibrated lower phase. The desired lower phases were then pooled and dried with a stream of N2. The purification yielded about 1.0 mg of 1-dephospho-lipid IVA.
The 1H NMR analysis (2D COSY) of the final compound
redissolved in 600 µl of CDCl3, CD3OD,
D2O (2:3:1, v/v) and analyzed under the conditions
described previously (42) confirmed its identity as 1-dephospho-lipid
IVA (supplementary figure). The spectrum of the purified
1-dephospho-lipid IVA, which is referenced against internal
tetramethylsilane, indicates that about 90% of the sample adopts the
-anomeric configuration in the proximal glucosamine unit, but the
-anomer and its cross-peaks are also detected. Following NMR
spectroscopy, the 1-dephospho-lipid IVA was dried down
under N2 and resuspended in 750 µl of 50 mM
MOPS, pH 7.0, to make a 1 mM aqueous dispersion. This was
stored at
80 °C and subjected to brief sonic irradiation prior to
use in assays.
Preparation of [4'-32P]1-dephospho-lipid IVA-- The starting material [4'-32P]lipid IVA was synthesized enzymatically as previously described (43) and then was converted into [4'-32P]1-dephospho-lipid IVA by hydrolysis in 0.1 M HCl. Typically, a sample containing ~80-100 µCi of [4'-32P]lipid IVA was re-suspended in 150 µl of 0.1 M HCl. Following sonic irradiation for 2 min in a bath apparatus, the mixture was placed in a 100 °C heat block for 15 min. After cooling, the hydrolyzed material was centrifuged briefly and then spotted onto a 20 × 20-cm Silica gel plate. Preparative TLC was carried out as described above for the nonradiolabeled lipid IVA. After drying, the product was detected by a brief autoradiography and recovered by extraction from the silica chips (see above). The final substrate was typically dispersed in 50 mM HEPES, pH 7.5, such that the working stock solution contained ~50,000 cpm/µl.
Assay Conditions for Measuring the Conversion of [14C]B to [14C]D-1-- The standard reaction mixture (10 µl) contained 10 µM [14C]B (~500 cpm/reaction), 0.5-1.0 mg/ml membrane protein, 0.1% Triton X-100, 1 mM MgCl2, and 50 mM MES buffer, pH 6.5, unless otherwise indicated. The reactions were incubated under ambient conditions at 30 °C and terminated at the indicated times by spotting 4-µl samples onto a 20 × 20-cm silica gel TLC plate. The spots were dried for 30 min with a cold air stream, and the plate was then developed in the solvent CHCl3, MeOH, H2O/pyridine (40:25:4:2, v/v). The remaining substrate and product(s) were detected with a Molecular Dynamics Storm PhosphorImager equipped with ImageQuant software. Enzyme specific activity (usually expressed as nmol/min/mg) was calculated based on the percentage of conversion of substrate to product(s).
Assay Conditions for Detecting the Oxidation of [4'-32P]1-Dephospho-lipid IVA-- The standard reaction mixture (10 µl) contained 5-10 µM [4'-32P]1-dephospho-lipid IVA (~5000 cpm/tube), 0.5-1 mg/ml membrane protein, 1 mM MgCl2, and 50 mM MOPS, pH 7.0, to partially suppress 4' phosphatase activity. After incubation at 30 °C for the indicated times, the reactions were stopped by spotting 4-µl samples onto a 20 × 20-cm silica gel TLC plate. The plate was developed with the solvent CHCl3, pyridine, 88% formic acid, H2O (50:50:16:5, v/v) and imaged as described above.
Isolation of the Product Generated from Component B in Vitro by
R. leguminosarum Membranes--
A 7.3-ml reaction mixture containing
50 µM R. etli component B (24, 25), 50 mM MES, pH 6.5, 1 mM MgCl2, 0.1%
Triton-X-100, and 0.5 mg/ml R. leguminosarum 3855 membranes
was incubated overnight (17.5 h) at 30 °C. TLC analysis of the
reaction at the initial and final time points indicated complete
conversion of B to a compound migrating with a component D-1 standard.
Half of the reaction mixture (contained in a sterile 16 × 125-mm
glass tube) was transferred to an identical glass tube. Each half was
then converted into a two-phase Bligh-Dyer mixture by the addition of
CHCl3 (4 ml) and MeOH (4 ml) to each tube. After mixing,
the phases were separated by centrifugation for 10 min at room
temperature in a clinical centrifuge. The lower phase was recovered and
transferred to a clean glass tube. After two extractions of the upper
phases with pre-equilibrated lower phases, all of the resulting lower phases were pooled, dried under N2, and stored at
20 °C.
A 1.5-ml DEAE-cellulose column (Whatman DE-52) (24) was used to purify
the lipids in the reaction mixture. The dried reaction product was
dissolved in 1.6 ml of CHCl3, MeOH, H2O (2:3:1,
v/v). After equilibration of the DEAE column as the acetate form in CHCl3, MeOH, H2O (2:3:1, v/v), the sample was
loaded and the flow through was collected as one fraction. The column
was then washed with 1.5 ml of CHCl3, MeOH, H2O
(2:3:1, v/v), which was collected as one fraction. Next, the column was
washed successively with the following solvent mixtures: 8 ml of
CHCl3, MeOH, 30 mM aqueous NH4Ac
(2:3:1, v/v), 8 ml of CHCl3, MeOH, 60 mM
aqueous NH4Ac (2:3:1, v/v), 8 ml of CHCl3,
MeOH, 120 mM aqueous NH4Ac (2:3:1, v/v), 6 ml
of CHCl3, MeOH, 240 mM aqueous
NH4Ac (2:3:1, v/v), and finally 6 ml of CHCl3,
MeOH, 500 mM aqueous NH4Ac (2:3:1, v/v). 1-ml
fractions were collected throughout the elution. Silica gel TLC
analysis of the fractions was done with the solvent CHCl3,
pyridine, 88% formic acid, MeOH, H2O (60:35:10:5:2, v/v).
The in vitro product emerged in the late 60 mM
NH4Ac wash, in accordance with the behavior of R. etli lipid A component D-1 (24). Lipids in fractions from the
flow-through, 30 mM NH4Ac, and 60 mM NH4Ac washes were pooled separately and
recovered by two-phase Bligh-Dyer partitioning, as discussed above for
the preparation of 1-dephospho-lipid IVA. The lower phases
for each pool were dried down with a stream of N2 and
stored at 20 °C.
Isolation of the Products Generated in Vitro from 1-Dephospho-lipid IVA by R. leguminosarum Membranes-- In a sterile 16 × 125-mm glass tube equipped with Teflon-lined cap, 100 µM 1-dephospho-lipid IVA was incubated with 0.36 mg/ml R. leguminosarum 3855 membranes in a buffer containing 50 mM MOPS, pH 7.0, 0.1% Triton X-100, and 1 mM MgCl2. The final reaction volume was 7 ml. A parallel reaction in a 1.5-ml microcentrifuge tube containing 96 µl of the above reaction mixture and 4 µl of [4'-32P]lipid IVA ~(50,000 cpm/µl) was carried out to monitor product formation with a PhosphorImager.
After 14 h at 30 °C, half of the nonradioactive reaction
mixture was transferred to a second identical glass tube.
CHCl3 (4 ml), MeOH (4 ml), and 0.1 M HCl (360 µl) were added to each of the two tubes, which were mixed and
centrifuged for 10 min in a clinical centrifuge. After two more
extractions of the upper phase with pre-equilibrated lower phase, all
of the lower phases were pooled in a clean glass tube. Two drops of
pyridine were added before the pooled lower phases were dried with
N2 and stored at 20 °C.
The sample was redissolved in 5 ml of CHCl3, MeOH,
H2O (2:3:1 v/v) and loaded onto a 1.0-ml DEAE-cellulose
column (Whatman DE-52) previously equilibrated as the acetate form in
CHCl3, MeOH, H2O (2:3:1, v/v) (24). The column
was washed with 1 ml of CHCl3, MeOH, H2O
(2:3:1, v/v). Elution was done stepwise with the following solvent
mixtures: 6 ml of CHCl3, MeOH, 30 mM aqueous
NH4Ac (2:3:1, v/v), 6 ml of CHCl3, MeOH, 60 mM aqueous NH4Ac (2:3:1, v/v), 5 ml of
CHCl3, MeOH, 85 mM aqueous NH4Ac
(2:3:1, v/v), 6 ml of CHCl3, MeOH, 120 mM
aqueous NH4Ac (2:3:1, v/v), 5 ml of CHCl3,
MeOH, 240 mM aqueous NH4Ac (2:3:1, v/v), and 5 ml of CHCl3, MeOH, 500 mM aqueous
NH4Ac (2:3:1, v/v). 0.5-ml fractions were collected from the 30 mM to the 120 mM NH4Ac
washes, and 1-ml fractions were collected for the subsequent steps. Ten
µl of each fraction was analyzed using silica gel TLC plates
developed in CHCl3, pyridine, 88% formic acid,
H2O (50:50:16:5, v/v). Fractions from each elution step
were pooled separately and recovered by two-phase Bligh-Dyer partitioning as described above for 1-dephospho-lipid IVA.
The desired lower phases were dried down, and the lipids were stored at
20 °C prior to further analysis.
Mass Spectrometry--
Matrix-assisted laser
desorption ionization time-of-flight (MALDI/TOF) mass spectra were
acquired on a Kompact MALDI 4 from Kratos Analytical (Manchester, UK),
equipped with a nitrogen laser (337 nm), 20 kV extraction voltage, and
time delayed extraction. The samples were prepared for MALDI/TOF
analysis by depositing 0.3 µl of the lipid sample dissolved in
chloroform/methanol (4:1, v/v), followed by 0.3 µl of a saturated
solution of 2,5-dihydroxybenzoic acid in 50% acetonitrile as the
matrix. The samples were left to dry at room temperature before the
spectra were acquired in both the positive and negative ion linear
modes. Each spectrum was the average of 50 laser shots.
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RESULTS |
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Conversion of [14C]B to [14C]D-1 by R. leguminosarum and R. etli Membranes--
As shown in Fig.
2, the membranes of R. leguminosarum 3855 efficiently convert [14C]B to a
substance migrating like [14C]D-1 in the presence of
Triton X-100. The cytosolic fraction is completely inactive (Fig. 2),
and it does not further stimulate the activity present in membranes
(not shown). Other nonionic detergents such as Nonidet P-40 can
substitute for Triton X-100, but ionic detergents, like SDS or LDAO,
are inhibitory (not shown). Product formation is nearly linear with
time for about 30 min at 0.25 mg/ml membrane protein (Fig.
3). The enzyme is inhibited by addition
of 5 mM EDTA to the assay mixture (Fig.
4A, lane 2a). However, nearly complete reactivation of the EDTA-inhibited enzyme is
observed upon addition of excess (10 mM) MgCl2
and incubation for another 20 min (Fig. 4B, lane
7). Co2+, Ni2+, and Mn2+ are
comparable with Mg2+ in reactivating the EDTA-treated
enzyme (Fig. 4B). Ca2+, Zn2+,
Cu2+, and Fe2+ are minimally effective, but
Mo2+ is inactive (Fig. 4B). The chelators
dipyridyl and o-phenanthroline did not inhibit the reaction
when added at 10 mM.
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The membranes of other strains of R. leguminosarum or
R. etli catalyze the conversion of [14C]B to
[14C]D-1 at rates that are comparable with those seen
with membranes of R. leguminosarum 3855 (Fig.
5). The membranes of E. coli
and S. meliloti 1021, which make lipid A species that are
fully phosphorylated and do not contain 2-aminogluconate, are unable to
metabolize [14C]B to [14C]D-1 (Fig. 5).
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MALDI/TOF Mass Spectrometry of the D-1-like Substance Synthesized in Vitro-- The TLC method (Figs. 2-5) used to monitor the conversion of [14C]B to [14C]D-1 by R. leguminosarum membranes strongly suggests that the product accumulating with time is authentic D-1 (Fig. 1), given its migration with a D-1 standard isolated from R. etli (24, 25). In addition, the D-1-like material synthesized in vitro has the same affinity as authentic D-1 for DEAE-cellulose (24), from which it elutes with CHCl3, MeOH, 120 mM aqueous NH4Ac (2:3:1, v/v) (data not shown). In contrast, the substrate [14C]B elutes with CHCl3, MeOH, 30 mM aqueous NH4Ac (2:3:1, v/v) because it is less negatively charged (24).
A scaled-up, nonradioactive enzymatic reaction mixture was used to
prepare D-1 for mass spectrometry. Following purification on a
DEAE-cellulose column, MALDI/TOF mass spectra of the remaining substrate B (Fig. 6, lower
panel) and of the D-1-like material (Fig. 6, upper
panel) were recorded in the positive reflectron mode. As noted in
Fig. 1, both the substrate and the product are mixtures of related
lipid species differing in acyl chain length by two methylene groups
(i.e. 28 atomic mass units) on the proximal unit (24, 25).
The observed peaks at m/z 1980.1 and 2008.1 (Fig.
6, lower panel) may be interpreted as the monoisotopic [M + Na]+ ions for the two predominant acyl chain lengths
present in B (Fig. 1) (24). The putative D-1 generated in
vitro by R. leguminosarum membrane gives strong peaks
at m/z 1996.5 and 2024.5 (Fig. 6, upper
panel), consistent with the incorporation of a single oxygen atom
by oxidation of the proximal glucosamine unit to the 2-aminogluconate residue (24). As with B, both signals may be interpreted as the
monoisotopic [M + Na]+ ions arising from the two
predominant acyl chain lengths present in D-1 (Fig. 1). The
distributions of the additional peaks at progressively higher
m/z in each of the clusters shown in Fig. 6 are
consistent with the molecular weights of B and D-1 (Fig. 1) and are in
agreement with previous studies of the natural products isolated from
R. etli CE3 using conventional MALDI/TOF mass spectrometry (24). The additional microheterogeneity of B and D-1 because of the
partial substitution with -hydroxybutyrate (Fig. 1) (24) is not seen
in the region of the spectrum shown in Fig. 6.
|
Use of [4'-32P]1-Dephospho-lipid IVA as
an Alternative Substrate for Demonstrating Oxidation of the Proximal
Glucosamine Unit--
The E. coli lipid A precursor
[4'-32P]lipid IVA, which is a tetra-acylated
disaccharide of glucosamine that is phosphorylated at positions 1 and
4' (36, 44), is not metabolized appreciably by R. leguminosarum membranes under the oxidase assay conditions (Fig.
7A, lower arrow). A
1-phosphatase is present in R. leguminosarum that converts
R. leguminosarum lipid A precursors to their
1-dephosphorylated derivatives (45), but this activity is barely
detectible with [4'-32P]lipid IVA as the
substrate under the conditions employed. However, when
[4'-32P]1-dephospho-lipid IVA (Fig.
8 and supplementary figure) is prepared by chemical hydrolysis of lipid IVA with 0.1 M
HCl at 100 °C and used as a substrate under the oxidase assay
conditions, it is metabolized to several new products (Fig.
7B). These compounds migrate slightly faster than the
substrate in this solvent system (Fig. 7B). Almost all of
the 1-dephospho-lipid IVA is consumed after overnight
incubation at 30 °C, as shown by mass spectrometry (see below). The
same lipid A oxidizing enzyme that converts B to D-1 is responsible for
the metabolism of 1-dephospho-lipid IVA by R. leguminosarum membranes (89).
|
|
Negative Ion MALDI/TOF Mass Spectrometry of the Products
Made by R. leguminosarum Membranes from 1-Dephospho-lipid
IVA--
The conversion of 1-dephospho-lipid
IVA to its putative oxidation products (Figs. 7 and 8) was
scaled up to permit mass spectrometry, as described under
"Experimental Procedures." Fig. 9
shows the MALDI/TOF spectra in the negative mode of the
1-dephospho-lipid IVA substrate, its putative lactone
derivative, and the proposed 2-aminogluconate product (Fig. 8). The
latter two compounds elute from DEAE-cellulose at about
CHCl3, MeOH, 60 mM aqueous NH4Ac (2:3:1, v/v) and CHCl3, MeOH, 120 mM aqueous
NH4Ac (2:3:1, v/v), respectively.
|
The substrate 1-dephospho-lipid IVA (Fig. 9A)
shows the expected peak at m/z 1324.8, which is
interpreted as [M H]
. The proposed
2-aminogluconate product (Fig. 9C) displays an intense peak
at m/z 1341.2, consistent with [M
H]
of a compound that has gained an oxygen atom (Fig.
8). The peak at m/z 928.9 (Fig. 9C)
could not be assigned and might be an impurity carried over from the
R. leguminosarum membranes used as the enzyme source.
The negative mode MALDI/TOF mass spectrum of the second (less abundant)
product derived from 1-dephospho-lipid IVA, which emerges
from DEAE-cellulose with the 60 mM NH4Ac wash,
is shown in Fig. 9B. An intense signal is observed at
m/z 1078.9. The putative lactone derivative of
1-dephospho-lipid IVA (Fig. 8) has a molecular weight of
1323.7 and should give rise to a peak at m/z
1322.7 in the negative mode. However, such lactones might undergo
elimination of the fatty acid moiety at position 3 (24, 25). In the
case of the putative lactone derived from 1-dephospho-lipid
IVA, the elimination of hydroxymyristic acid would generate
an /
-unsaturated lactone (Fig. 8) with a molecular weight of
1079.3. Consequently, the major peak seen at m/z
1078.9 in Fig. 9B could be interpreted as [M
H]
derived from the lactone elimination product (Fig.
8). We do not believe that the putative elimination product forms prior to mass spectrometry, because it would migrate more slowly during TLC
than 1-dephospho-lipid IVA, which is not observed (Fig.
7B).
A small amount of residual substrate is evident in the sample shown in Fig. 9B, consistent with its charge (Fig. 8), as judged by the peak at m/z 1325.2. As in Fig. 9C, the peak at m/z 928.9 (Fig. 9B) is not assigned and may represent an impurity.
Positive Ion MALDI/TOF Mass Spectrometry of Products
Made by R. leguminosarum Membranes from 1-Dephospho-lipid
IVA--
The positive ion MALDI/TOF mass spectra (Fig.
10) of the same three substances are
consistent with the interpretation of their negative mode spectra (Fig.
9). Most significantly, the positive mode spectra reveal an intense
B
|
Outer Membrane Localization of the Lipid A Oxidase of R. leguminosarum--
In previous studies, we have described methods for
separating inner and outer membranes of various Rhizobium
strains using isopycnic sucrose gradient centrifugation (46). As in
E. coli, the heavier (outer) membrane fraction is
characterized by its phospholipase A activity, whereas the lighter
(inner) membranes are detected using NADH oxidase. Remarkably, the
lipid A oxidase activity, which was measured by following the
conversion of [14C]B to [14C]D-1, is
recovered almost entirely with the outer membrane fragments of R. leguminosarum 3855 (Fig. 11). The
same profile was obtained when 1-dephospho-lipid IVA was
used as the substrate (not shown). Among the various R. etli
or R. leguminosarum lipid A enzymes studied to date
(45-51), only the oxidase is found in the outer membrane. The outer
membrane localization of the oxidase therefore demonstrates that the
formation of 2-aminogluconate represents a late modification in
the maturation of R. leguminosarum lipid A.
|
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DISCUSSION |
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R. leguminosarum and R. etli are the only organisms known to synthesize 2-aminogluconate, an oxidized derivative of D-glucosamine (23-25, 52). This material substitutes for the proximal glucosamine unit that is usually present in lipid A (Fig. 1) (23-25, 52). In re-evaluating the microheterogeneity and structures of R. leguminosarum and R. etli lipid A, Que et al. (24, 25, 28) discovered that about one-third of the lipid A isolated from these organisms does in fact contain the conventional glucosamine disaccharide backbone found in most other Gram-negative bacteria (Fig. 1, structure B). In earlier studies by Bhat et al. (23), B had been overlooked, because the lipid A released from lipopolysaccharide by acid hydrolysis was not further purified. Given that B lacks the anomeric phosphate residue (24, 25), it might be the immediate precursor of component D-1 (Fig. 1). We have now developed a quantitative assay for following the conversion of B to D-1 using membranes of R. leguminosarum or R. etli (Figs. 2-5) and have demonstrated the presence of an additional oxygen atom in the D-1 product generated in vitro from B. We also report the model substrate, 1-dephospho-lipid IVA (Figs. 7 and 8), with which to characterize the enzyme.
A remarkable feature of the oxidase is its outer membrane localization. This finding suggests that the formation of 2-aminogluconate occurs as a late modification of the lipid A molecule. All of the early conserved reactions of lipid A biosynthesis in R. leguminosarum and R. etli, as well as the 4' and 1 phosphatases that are unique to these organisms, appear to be associated with the inner membrane (45-51).2
Very few outer membrane enzymes have ever been described. All of the known outer membrane enzymes are lipases (35, 53), acyltransferases (54, 55), or proteases (56). The outer membrane localization of the enzyme that generates D-1 from B suggests that it is a novel kind of oxidase. Oxygen is indeed required for the reaction, as demonstrated in the accompanying manuscript. However, it is not yet possible to show stoichiometric formation of H2O2 and D-1 from O2 and B when using whole membrane preparations as the enzyme source (see below).
The ability of the oxidase to utilize 1-dephospho-lipid IVA (Figs. 7 and 8) as a model substrate demonstrates that it does not require the 2' acyloxyacyl or the 4' galacturonic acid residues characteristic of R. leguminosarum or R. etli lipid A (Fig. 1) for catalysis. However, the oxidase does require a free hydroxyl group at the anomeric position of the proximal lipid A unit, as shown by its inability to utilize lipid IVA (Fig. 7A). The oxidase also prefers a substrate with an acyl chain at position 3, given its relatively slow oxidation of component C (Fig. 1) compared with B (data not shown).
Two plausible routes for the oxidation of the proximal glucosamine unit
are shown in Fig. 8. In the lactone pathway (Fig. 8, reactions
2 and 3), oxidation occurs before ring opening, and a
lactone intermediate is formed that could be hydrolyzed by a separate
lactonase to generate the 2-aminogluconate moiety. Alternatively, the
same protein might catalyze both the oxidation and lactone hydrolysis,
or the latter step might even be nonenzymatic. In any case, an
analogous lactone is synthesized by glucose oxidase and glucose
dehydrogenase. Glucose oxidase from Aspergillus niger (57)
is a soluble iron-dependent flavoenzyme (58) that converts glucose and oxygen to -gluconolactone and
H2O2. On the other hand, the glucose
dehydrogenases are generally PQQ-dependent enzymes that likewise convert glucose to gluconolactone (59). Depending on the
microorganism, glucose dehydrogenase can be membrane-bound or soluble
and requires either Mg2+ or Ca2+ (59).
Membrane-bound glucose dehydrogenase has been isolated from a wide
range of bacteria (59). Its active site is presumed to be located on
the periplasmic surface of the inner membrane, and the electron
acceptor is ubiquinone (60, 61). A conserved region near the N
terminus, predicted to consist of five membrane-spanning helices, is
thought to be important for ubiquinone binding (62).
Although the membrane-bound glucose dehydrogenases oxidize various monosaccharides, they cannot process disaccharides. In contrast, an atypical soluble glucose dehydrogenase found only in Acinetobacter calcoaceticus (63, 64) can oxidize both mono- and disaccharides. Interestingly, this periplasmic enzyme is rather different from the membrane-bound glucose dehydrogenases, not only with regard to its sugar substrate specificity but also in its preference for electron acceptors. It does not utilize ubiquinone (61) but instead slowly reduces a soluble cytochrome b that does not appear to interact with the electron transport chain (65). Furthermore, the periplasmic dehydrogenase shares very little amino acid sequence similarity with the membrane-bound enzymes (66), and it lacks the conserved 11-residue tryptophan docking motif found in most other PQQ-containing dehydrogenases (67).
Another enzyme that oxidizes the C1 of a hexose moiety is cellobiose dehydrogenase, which is found in the lignin-degrading white rot fungi Phanerochaete chrysosporium (68-72) and Humicola insolens (73). Cellobiose dehydrogenase is an extracellular enzyme that oxidizes various di- and oligosaccharides and can utilize either Fe3+, O2, or various organic molecules as electron acceptors (74, 75). As shown in the accompanying manuscript, the lipid A oxidase of R. leguminosarum does not share significant sequence homology with any of the above enzymes.
Another alternative for 2-aminogluconate formation might involve ring opening of the proximal lipid A unit, followed by oxidation the aldehyde (Fig. 8, reaction 1). This possibility is reminiscent of the mechanism proposed for D-xylose isomerase (76) (also known as D-glucose isomerase), which interconverts a broad spectrum of aldoses and ketoses (77). X-ray data suggest the presence of an extended open chain sugar substrate in the enzyme active site (78, 79). Mg+2 is believed to be the sole cofactor needed by the enzyme in vivo (80), but other divalent cations can be substituted in vitro.
Given the outer membrane localization of the lipid A oxidase, we consider it unlikely that its enzymatic mechanism involves pyridine or flavin nucleotides. This idea is supported by the observation that there is no stimulatory effect on the rate of conversion of B to D-1 by addition of exogenous NAD, NADP, FAD, FMN, ubiquinone, or cytochrome c (data not shown). Furthermore, the reaction is not inhibited by cyanide. A mechanism involving PQQ deserves consideration in view of the inner membrane or periplasmic localization of the PQQ-dependent glucose dehydrogenases discussed above, but again no stimulation of B to D-1 conversion was seen with added PQQ (data not shown). At present, the only clue to the mechanism of the oxidation is the inhibition by added EDTA and the reactivation with excess Mg2+ and some other divalent cations, especially Co2+, Ni2+, and Mn2+ (Fig. 4). Although a direct involvement of Mg2+ in substrate binding, catalysis, or maintenance of tertiary structure is certainly a possibility, we cannot exclude the alternative that the enzyme requires a redoxactive heavy metal, which is removed by EDTA and then transferred back in the presence of excess Mg2+ or other divalent cations (Fig. 4).
A mechanism in which electrons are transferred from B to molecular oxygen in the outer membrane without the involvement of the inner membrane electron transport chain seems attractive based upon the available data. This hypothesis predicts that H2O2 would be formed as a by-product. Attempts to demonstrate stoichiometric formation of H2O2 with the Amplex Red detection kit (Molecular Probes) during the conversion of 50 µM B to D-1 have not been successful so far (data not shown). However, the membrane preparations used as the enzyme source rapidly consume 50 µM H2O2 added exogenously (data not shown). Purification of the oxidase to homogeneity, as well as structural and mechanistic studies, will therefore be required to identify the intermediates and by-products of the conversion of B to D-1.
Although the occurrence of 2-aminogluconate is limited to a few
Gram-negative bacteria involved in nitrogen fixation (23-25), an
emerging general theme in lipid A biogenesis is that specific covalent
modifications may occur within the outer membrane (31, 54, 55). In some
cases, these outer membrane modifications are subject to exquisite
transcriptional regulation (31, 54). For instance, the lipid A
acyltransferase PagP of E. coli and S. typhimurium is regulated by the PhoP/PhoQ two-component system (54), which is induced during growth on low concentrations of Mg2+ (81, 82). PagP is a typical outer membrane protein and
is synthesized with a signal sequence that is cleaved during export (54). Recent structural NMR studies of PagP (55) have revealed that its
active site faces the outside. It will be very interesting to determine
the structure of the oxidase and establish whether or not it is
regulated, especially during the differentiation of bacteroids to form
nitrogen-fixing nodules. Mutants lacking the lipid A oxidase will be
necessary to study the function of the 2-aminogluconate residue. The
identification of the structural gene encoding the oxidase, described
in the accompanying article (89), should facilitate such further
biological and chemical studies.
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ACKNOWLEDGEMENTS |
---|
We thank Dr. Tony Ribeiro at the Duke University NMR Center for help with instrumentation and Suzanne Kalb of the Johns Hopkins University for recalibration of the MALDI/TOF spectra. We thank Dr. Tom Krick of the University of Minnesota Mass Spectrometry Center, College of Biological Sciences (St. Paul campus) for carrying out the MALDI/TOF experiments in the reflectron mode.
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FOOTNOTES |
---|
* This work was supported by National Institutes of Health Grants R37-GM-51796 (to C. R. H. R.) and GM-54882 (to R. J. C.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
The on-line version of this article (available at
http://www.jbc.org) contains a supplemental figure.
¶ Present address: Ciphergen Biosystems, Inc., Fremont, CA 94555.
To whom correspondence should be addressed. Tel.:
919-684-5326; Fax: 919-684-8885; E-mail: raetz@biochem.duke.edu.
Published, JBC Papers in Press, January 15, 2003, DOI 10.1074/jbc.M300378200
2 M. J. Karbarz and C. R. H. Raetz, manuscript in preparation.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: LPS, lipopolysaccharide; 2-aminogluconate, 2-amino-2-deoxy-gluconate; MES, 2-(N-morpholino)-ethanesulfonic acid; MOPS, 3-(N-morpholino)-propanesulfonic acid; MALDI, matrix-assisted laser desorption ionization; TOF, time-of-flight; PQQ, pyrroloquinoline quinone.
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