From the Department of Biochemistry, Duke University
Medical Center, Durham, North Carolina 27710 and the ¶ Middle
Atlantic Mass Spectrometry Laboratory, Department of Pharmacology and
Molecular Sciences, The Johns Hopkins University of School of
Medicine, Baltimore, Maryland 21205-2185
Received for publication, January 13, 2003, and in revised form, January 15, 2003
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ABSTRACT |
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An unusual feature of the lipid A from the plant
endosymbionts Rhizobium etli and Rhizobium
leguminosarum is the presence of a proximal sugar unit consisting
of a 2-amino-2-deoxy-gluconate moiety in place of glucosamine. An outer
membrane oxidase that generates the 2-amino-2-deoxy-gluconate unit from
a glucosamine-containing precursor is present in membranes of R. leguminosarum and R. etli but not in S. meliloti or Escherichia coli. We now report the identification of a hybrid cosmid that directs the overexpression of
this activity by screening 1800 lysates of individual colonies of a
R. leguminosarum 3841 genomic DNA library in the host
strain R. etli CE3. Two cosmids (p1S11D and p1U12G) were
identified in this manner and transferred into S. meliloti,
in which they also directed the expression of oxidase activity in the
absence of any chromosomal background. Subcloning and sequencing of the
oxidase gene on a 6.5-kb fragment derived from the ~20-kb insert in
p1S11D revealed that the enzyme is encoded by a gene (lpxQ)
that specifies a protein of 224 amino acid residues with a putative
signal sequence cleavage site at position 28. Heterologous expression
of lpxQ using the T7lac promoter system in
E. coli resulted in the production of catalytically active
oxidase that was localized in the outer membrane. A new outer membrane
protein of the size expected for LpxQ was present in this construct and
was subjected to microsequencing to confirm its identity and the site
of signal peptide cleavage. LpxQ expressed in E. coli
generates the same products as seen in R. leguminosarum
membranes. LpxQ is dependent on O2 for activity, as
demonstrated by inhibition of the reaction under strictly anaerobic conditions. An ortholog of LpxQ is present in the genome of
Agrobacterium tumefaciens, as shown by heterologous
expression of oxidase activity in E. coli.
As demonstrated in the accompanying article (1), the outer
membranes of Rhizobium leguminosarum and Rhizobium
etli contain an unusual oxidase that converts the proximal
glucosamine unit of 1-dephosphorylated lipid A (or related molecules)
to a novel 2-aminogluconate moiety. The membranes of
Sinorhizobium meliloti and Escherichia coli do
not normally contain such an oxidase activity. Although the function of
this unusual covalent modification of lipid A is unknown (2-4), the
existence of an oxidative enzyme in the outer membrane of a
Gram-negative bacterium is without precedent. The few outer membrane
enzymes described to date are all phospholipases (5, 6),
acyltransferases (7, 8), or proteases (9). Other characterized outer
membrane proteins function either as porins or specialized transporters
(10-12). The presence of the oxidase in outer membranes of certain
strains of Rhizobium suggests that lipid A oxidation (1),
when it occurs, is a late event in lipopolysaccharide assembly.
Given the considerable progress that has recently been made with the
structural biology of outer membrane proteins by x-ray crystallography
(12) and NMR spectroscopy (8, 13) and the great interest in lipid A
modifications in the context of microbial pathogenesis (14-16), we now
report the expression cloning of a novel R. leguminosarum
gene, designated lpxQ, that encodes the oxidase. Hydropathy
analysis predicts that LpxQ is a typical outer membrane protein of 224 amino acid residues with a leader sequence that is cleaved during
export. The R. leguminosarum oxidase is properly localized
to the outer membrane when expressed in E. coli behind the
T7lac promoter. The only other bacterial genomes that
contain clear-cut orthologs of LpxQ are those of two
Agrobacterium tumefaciens strains, the sequences of which
were recently reported (17, 18). Although 2-aminogluconate has not been
described as a component of lipid A in A. tumefaciens, we
show that A. tumefaciens LpxQ is in fact fully active as a
lipid A oxidase when overexpressed in E. coli. The
availability of the lpxQ genes of R. leguminosarum, R. etli, and A. tumefaciens
should enable purification to homogeneity and mechanistic studies of
the oxidase and should also facilitate genetic studies of
2-aminogluconate function during plant infection and symbiosis.
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, and 14C-labeled
component B was prepared as described in the accompanying article
(1).
Bacterial Growth Conditions--
Briefly, 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. Unless otherwise indicated, R. etli CE3 (19, 20) and S. meliloti 1021 (21, 22) 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 generally grown at 37 °C in LB broth (23) 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 plasmids and bacterial strains used in this study.
Cell-free extracts and washed membranes were prepared as described in
the accompanying article (1).
Quantitative Assay for Measuring the Conversion of
[14C]B to [14C]D-1--
As described
fully in the accompanying article (1), the standard reaction mixture
(10 µl) contained (unless otherwise indicated) 10 µM
[14C]B (~500 cpm/tube), 0.5-1.0 mg/ml membrane
protein, 0.1% Triton X-100, 1 mM MgCl2, and 50 mM MES1 buffer,
pH 6.5. The reactions were incubated under aerobic conditions at
30 °C and terminated at the indicated times by spotting 4- or 5-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).
Anaerobic Assay Conditions--
For demonstrating oxygen
dependence, an assay system consisting of 25 µM
[14C]B, 50 mM MES buffer, pH 6.5, 0.1%
Triton X-100, 1 mM MgCl2, and 0.1 mg/ml
BLR(DE3)/pLysS/pQN235 membranes was set up in an anaerobic chamber. The
reactions were started by the addition of membranes as the enzyme
source after equilibration of all tubes in the absence of oxygen for
~30 min. To remove traces of oxygen, glucose oxidase (20 µg/ml),
catalase (2 µg/ml), and glucose (0.1%) were included in some cases,
as indicated. After 30, 60, or 90 min, portions of the reaction
mixtures were spotted onto a silica gel TLC plate, which was developed
as described above. A parallel set of reaction mixtures was assayed
simultaneously under ambient aerobic conditions.
Expression Cloning of the R. leguminosarum Lipid A Oxidase
Gene--
A genomic DNA library of the R. leguminosarum
strain 3841 was obtained from Dr. J. Downie of the John Innes Institute
(Norwich, UK). This library was constructed by the ligation of
~20-25-kb fragments of genomic DNA (24), generated by partial
EcoRI digestion, into the cosmid pLAFR-1 (25). E. coli 803 was employed as the host strain (26).
The library was transferred from E. coli 803 into R. etli CE3 by tri-parental mating (27, 28) with E. coli
MT616 as the helper (28). Three thousand individual R. etli
CE3 colonies harboring random fragments of the library were picked into
microtiter dishes containing (per well) 150 µl of TY medium with 20 µg/ml nalidixic acid, 200 µg/ml streptomycin, 12 µg/ml
tetracycline, and 10 mM CaCl2 and then were
grown at 30 °C with shaking at 225 rpm to stationary phase. The
cultures were stored at
The remaining 100 µl in each well of the microtiter plate was
centrifuged at 3,660 × g for 5 min at 4 °C. The
cell pellets were washed with 50 µl of 50 mM
HEPES, pH 7.5, and centrifuged as above. After decanting the
supernatant, the cell pellets were resuspended in 10 µl of lysis
buffer, which consists of 100 mg/ml lysozyme in 50 mM
HEPES, pH 7.5. The plates were sealed, and the contents were vigorously
mixed for a minute. Lysis was allowed to proceed at room temperature
for 30 min. The lysates were frozen by placing the plate into a
Three apparently positive pools of lysates were identified and
subjected to further analysis. Membranes from each of the nine cosmid-bearing strains contributing to the three positive pools were
prepared and assayed individually for overproduction of oxidase activity versus membranes of the control CE3 strain. Next,
the hybrid cosmids of the three strains that directed overexpression of
oxidase activity were isolated, transformed into E. coli
HB101, and then transferred into S. meliloti via
tri-parental mating (27, 28). Finally, membranes of S. meliloti cells harboring these cosmids were assayed for their
ability to express oxidase activity to verify the presence of the lipid
A oxidase gene on the cosmid inserts. Membranes of the S. meliloti 1021 host strain (22) used for this purpose (unlike CE3)
lack background oxidase activity.
Recombinant DNA Techniques--
Protocols for handling of DNA
samples were those of Sambrook et al. (29). Competent cells
of E. coli were prepared by the CaCl2 method
(29, 30). Cosmids and plasmids were purified using Qiagen DNA
purification kits. Double-stranded DNA sequencing was performed with an
ABI Prism 377 instrument at the Duke University DNA Analysis Facility.
Primers were purchased from Invitrogen. DNA sequence analysis and
searches for open reading frames were performed with GCG and ORF Finder
(31).
Subcloning and Sequencing of the DNA Inserts in Cosmids p1S11D
and p1U12G--
Digestion of the cosmid p1S11D with HindIII
generated fragments of ~0.2, 1.2, 3.5, 4, and 6.5 kb, as well as a
large band (~21 kb) corresponding to the vector pLAFR-1. The
insert-derived fragments were ligated into the shuttle vector pRK404a
digested with HindIII (32), and the desired constructs were
transferred into S. meliloti via tri-parental mating.
Membranes of S. meliloti cells harboring each of these
subclones were prepared and assayed for oxidase activity. Only one
hybrid plasmid, pQN210, which contains the 6.5-kb HindIII
fragment of p1S11D, directed the expression of oxidase activity. The
same 6.5-kb fragment was also present in the DNA insert of p1U12G and
was active when expressed on the hybrid plasmid pQN217 (Table I). The
6.5-kb HindIII fragment of pQN210 was sequenced using a
primer walking strategy (29). The DNA sequencing was facilitated by
subcloning of several EcoRI fragments derived from the
6.5-kb insert present in pQN210 into the vector pRK404a. None of these
subclones directed the expression of oxidase activity, given that an
EcoRI site is present in the oxidase structural gene
lpxQ (see below).
Construction of Plasmids That Express R. leguminosarum lpxQ
behind the lac or the T7lac Promoters--
The lpxQ gene
was cloned into several vectors, which were tested for their ability to
direct the overexpression of oxidase activity. The coding region of
lpxQ was amplified by PCR from pQN210 with various primers
to optimize restriction enzyme sites, translational start sites, or
translational stop sites.
For all of the primers, restriction enzyme sites are italicized, and a
G/C clamp is incorporated at their 5'-ends. The 5'-lpxQ/1 primer
(5'-CGCGCAAGCTTAGGAGGAATTTAAAATGACATATGCGCTGCGTTCTTCCG-3') was designed for ligation of lpxQ into lac
promoter-driven vectors, such as the shuttle vector pRK404a. This
primer also incorporates a HindIII site (in italics), a
ribosome-binding site (underlined sequence), and a translational spacer
element between the ribosome-binding site and the ATG start codon (33).
The sequences in bold type correspond to the N-terminal coding region
of lpxQ.
An internal NdeI site (CATAT9G) (in which the
superscript reflects the number of bases from the A of the ATG start
codon) is present in the coding sequence of lpxQ. The primer
5'-lpxQ/2
(5'-CGCGCCATATGACATAC9GCGCTGCGTTCTTCCG-3')
was designed to eliminate this internal site and to facilitate the
insertion of lpxQ into pET vectors behind the
T7lac promoter via an engineered NdeI site (in
italics). The internal NdeI site in the coding region of
lpxQ was removed by mutating T9 to
C9 in 5'-lpxQ/2. This base substitution does not
alter the amino acid sequence of LpxQ. Sequences in bold type
correspond to the N-terminal coding region of lpxQ.
The primer 3'-lpxQ/1
(5'-GCGCAAGCTTTTACACATATTCCCTGACGATAGCAGGC-3')
contains an engineered HindIII site (in italics). The underlined sequence corresponds to a region of DNA that is 96 bases
downstream of the termination codon of lpxQ. Lastly, The primer 3'-lpxQ/2
(5'-GCGCAAGCTTCCAGTGGAATGAAACGCCGACGTTGA-3') was designed for ligation of lpxQ into pET21b+, whereby a
protein is made with a His6 tag fused to the C terminus of
LpxQ. The sequence in bold type corresponds to the coding region at the
C-terminal end of lpxQ less the termination codon.
The following combinations of primers were used for PCRs and subsequent
ligations. The PCR product of 5'-lpxQ/1 and
3'-lpxQ/1 was digested with HindIII and ligated
into the shuttle vector pRK404a to yield pQN231. The lpxQ
gene amplified with primers 5'-lpxQ/2 and
3'-lpxQ/1 was digested with NdeI and
HindIII and then ligated into pET21a+, yielding pQN233. The
DNA generated with 5'-lpxQ/2 and 3'-lpxQ/2 was
digested with NdeI and HindIII and then ligated
into pET21b+, yielding pQN235.
A typical PCR mixture contained 100 ng of template DNA (pQN210), 1×
Pfu polymerase buffer (Stratagene), 10% Me2SO,
1% glycerol, 200 µM of each of the dNTPs, 125 ng of each
primer, and 2.5 units of Pfu polymerase (Stratagene) in a
total volume of 100 µl. The reaction was placed in a DNA thermal
cycler and subjected to 5 min of denaturation at 94 °C, followed by
five cycles consisting of denaturation (1 min, 95 °C), annealing (1 min, 55 °C), and extension (2 min, 72 °C). Full amplification was
then achieved with 25 cycles of the following: denaturation (1 min,
95 °C), annealing (1 min, 68 °C), and extension (2 min,
72 °C). After the 30th overall cycle, an additional
6-min extension at 72 °C was performed. The reaction was terminated
by cooling the tubes to 4 °C. In each case, the product was analyzed
on a 0.8% agarose gel, excised and purified using the Gene Clean II
gel DNA purification kit (Bio 101), digested with the appropriate
restriction enzymes as indicated above, and ligated into a vector that
had been similarly digested. The ligation mixtures that resulted in the
construction of pQN233 and pQN235 were first transformed into competent
E. coli XL-1Blue cells. The transformants were screened for
the desired insert by restriction enzyme digestion. The insert and
flanking regions of pQN233 were confirmed by DNA sequencing. The final recombinant plasmids were then transformed, as indicated below, into
competent E. coli cells of strain BLR(DE3)/pLysS or
Novablue(DE3) (Novagen) to evaluate the overexpression of LpxQ upon
induction of mid-log phase cells at 37 °C with 1 mM
IPTG.
The ligation mixture that yielded pQN231 was first transformed into
competent cells of E. coli HB101. The four candidate
plasmids containing the proper insert were then moved into S. meliloti 1021 by the tri-parental mating procedure. Of these four
hybrid plasmids, only two were found to direct the expression of LpxQ activity and were designated pQN231, presumably reflecting the proper
orientation of the lpxQ gene for expression behind the lac promoter. The membranes for assay of these constructs
were prepared as described in the accompanying article (1).
PCR Amplification and Cloning of an A. tumefaciens lpxQ
Ortholog--
The A. tumefaciens lpxQ gene was amplified by
PCR followed by ligation into the pET21b+ vector. The N-terminal primer
for the PCRs had the following sequence:
5'-CGCGCATATGTCGCGCCATACTTTACTTGTTTGC-3'. This primer was designed with a G/C clamp, an NdeI
restriction site (in italics) that overlaps with the initiation codon
ATG and the first 26 base pairs of the A. tumefaciens lpxQ
gene coding sequence (in bold type). The C-terminal primer was designed
with a G/C clamp, a BamHI site (in italics), and sequence
(in bold type) corresponding to a region that is 52 base pairs
downstream of the lpxQ TAA termination codon. The sequence
of the C-terminal primer was:
5'-GCGCGGATCCTGGAACTGCTGGACTGGGCTTATGG-3'.
A. tumefaciens C58 genomic DNA purchased from the American
Type Culture Collection (Manassas, VA) was used as the template. The
PCR mixture (100 µl) contained 250 ng of template DNA, 1× Pfu polymerase buffer (Stratagene), 10% Me2SO,
1% glycerol, 200 µM of each of the dNTPs, 125 ng of each
primer, and 2.5 units of Pfu polymerase (Stratagene). The
reaction was placed in a DNA thermal cycler and subjected to a 5-min
denaturation at 94 °C, followed by five cycles consisting of
denaturation (1 min, 95 °C), annealing (1 min, 55 °C), and
extension (2 min, 72 °C). Full amplification was then achieved with
25 cycles of the following: denaturation (1 min, 95 °C), annealing
(1 min, 65 °C), and extension (2 min, 72 °C). After the
30th overall cycle, an additional 6-min extension at
72 °C was performed. The PCR was terminated by cooling the tubes to
4 °C.
The PCR product was analyzed on a 0.9% agarose gel and was
gel-purified with the GeneClean DNA system (see above). The purified lpxQ gene, and the vector were both digested with
NdeI and BamHI and then ligated at 16 °C
overnight. The ligation mixture was transformed into competent E. coli XL-1Blue cells (Stratagene). Several ampicillin-resistant
colonies were picked, and the purified plasmids were screened for the
desired insert by digestion with NdeI and BamHI.
The construct harboring the A. tumefaciens lpxQ gene was
designated pQN240 and was transformed into BLR(DE3)/pLysS (Novagen) for
T7lac-directed overexpression.
Preparation of E. coli Membranes Containing A. tumefaciens LpxQ
for Assay--
For this purpose, a single colony of
BLR(DE3)/pLysS/pQN240 was used to inoculate 5 ml of LB broth containing
100 µg/ml ampicillin and 20 µg/ml chloramphenicol. After shaking at
225 rpm for 16 h at 37 °C, a portion of this overnight culture
was used to inoculate 50 ml of LB broth containing ampicillin and
chloramphenicol at an A600 of 0.01. The cells
were grown in shaking culture (225 rpm) at 37 °C until the
A600 had reached ~0.5, at which time 1 mM IPTG was added. After another 3 h of growth at
37 °C, the cells were harvested by centrifugation at 3660 × g for 15 min at 4 °C. The cells were washed once with 10 ml of 50 mM HEPES, pH 7.5, and again collected by
centrifugation. The cell pellet was then resuspended in 5 ml of 50 mM HEPES, pH 7.5, and stored at Subcellular Localization of LpxQ Expressed in E. coli--
The
subcellular localization of the R. leguminosarum LpxQ
oxidase expressed in E. coli Novablue(DE3)/pQN233 was
determined using a protocol similar to that described by Trent et
al. (34) and in the accompanying article (1). Briefly, a 500-ml
culture of Novablue(DE3)/pQN233 was grown with shaking (225 rpm) at
37 °C to A600 = 0.5 and then induced with 1 mM IPTG. After 3 h of further growth, the culture was
centrifuged at 6,000 × g for 10 min at 4 °C. The
cell pellet was resuspended in 10 ml of 50 mM HEPES, pH
7.5, and the cells were broken by two passages through a French press
cell at 10,000 p.s.i. Unbroken cells and large debris were removed by
centrifugation at 12,100 × g for 10 min at 4 °C.
The membranes were prepared and washed once as described in the
accompanying article (1). 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 layering on top of a seven-step isopycnic sucrose gradient Guy-Caffey et al. (35), the inner and outer membranes were separated by centrifugation in a Beckman SW41 swinging bucket rotor at 155,000 × g for 18 h at 4 °C. Fractions of 0.5 ml
were collected, and their protein content was determined using the
bicinchoninic method (36). The following volumes of each fraction were
used without dilution for the various enzyme assays: 5 µl for LpxQ,
50 µl for the NADH oxidase, and 10 µl for phospholipase A (34).
The inner and outer membranes were pooled separately. Each pool was
diluted 4-fold with 50 mM HEPES, pH 7.5, and the membrane fragments were collected by centrifugation at 149,000 × g for an hour. The membranes were resuspended in 0.15 ml of
50 mM HEPES, pH 7.5, and the protein concentrations were
determined by the bicinchoninic acid assay (36). Control membranes from
a 500-ml culture of Novablue(DE3) cells containing the empty
vector pET21a+ were prepared in parallel.
Protein Microsequencing of LpxQ--
Inner and outer membranes
of Novablue(DE3)/pET21a+ and Novablue(DE3)/pQN233 were analyzed by
SDS-PAGE on a Bio-Rad Protean II XI apparatus with a 12%
polyacrylamide, 1.5-mm thick gel (37). A band of the size expected for
LpxQ was observed in the outer membranes only. A piece of the gel
containing LpxQ was excised. The proteins were blotted onto an
Immobilon-P polyvinylidene fluoride membrane (Millipore) equilibrated
in 10 mM CAPS, pH 11, containing 10% methanol, at 15 V for
30 min, using a Bio-Rad Semi-Dry Transfer apparatus. To visualize the
transferred proteins, the blot was immersed in 0.1% Ponceau S in 1%
acetic acid for 1 min, followed by destaining with 1% acetic acid for
10 min. The band corresponding to LpxQ was excised, rinsed three times
with distilled water, and analyzed by high sensitivity protein
microsequencing on ABI model 492A Sequencer at the University of
Massachusetts Medical School Proteomic Mass Spectrometry Laboratory
(Worcester, MA).
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, as described in the accompanying article (1).
Expression Cloning of the Lipid A Oxidase of R. leguminosarum--
The gene encoding the lipid A oxidase was found by
assaying ~600 pools of three individual lysates of an R. leguminosarum genomic DNA library harbored in R. etli
CE3 for their ability to convert [14C]B to
[14C]D-1 (Fig.
1A). A basal level of oxidase
activity was present in all of the pools because of the chromosomal
copy of the oxidase gene present in the CE3 host. As shown in Fig.
1B, this background activity was quantified at about 11%
conversion of [14C]B to [14C]D-1 in 60 min
under the assay conditions employed. Occasional samples, such as the
one indicated by the arrow, derived from the pool of wells
10-12 from row D of plate 1S (Fig. 1B), catalyzed about
2.5-fold more rapid conversion of [14C]B to
[14C]D-1 than did the others. The lysates making up these
active pools were analyzed individually (data not shown). In this
manner, three positive cosmids (p1U12G, p1S11D, and p1E11D) capable of directing the overexpression of oxidase activity in CE3 were
identified. Because p1E11D and p1S11D contained exactly the same
inserts (data not shown), only p1S11D and p1U12G were further
characterized.
The active cosmids p1U12G and p1S11D were transferred via tri-parental
mating from an E. coli HB101 stock culture into S. meliloti 1021. The latter does not contain endogenous oxidase activity and lacks the 2-aminogluconate unit in its lipid A. S. meliloti expresses R. leguminosarum genes very
effectively from their native promoters, whereas E. coli
does not. As shown in Fig. 2, no
background oxidase activity is present in cell extracts or membranes of
the control strain S. meliloti/pLAFR-1 (lanes 2 and 5). In contrast, robust conversion of
[14C]B to [14C]D-1 is observed in both cell
extracts (lanes 3 and 4) and membranes (lanes 6 and 7) of S. meliloti/p1S11D
and S. meliloti/p1U12G. These results provide compelling
evidence for the presence of the lipid A oxidase gene on the inserts in
each of the above cosmids.
Subcloning of p1S11D Localizes the Oxidase Gene to a 6.5-kb HindIII
Fragment--
To determine the exact location of the oxidase gene, the
DNA inserts in both p1S11D and p1U12G were digested with
EcoRI and HindIII (Fig.
3), as well as with PstI (not
shown). The resulting DNA fragments were ligated into pRK404a and
transformed into E. coli HB101. The subclones obtained in
this manner were transferred into S. meliloti 1021 by
tri-parental mating (Fig. 3 and Table I). Upon assaying membranes of
the various constructs, only pQN210, which contains a ~6.5-kb
Hind III fragment (Table I) from p1S11D, directed expression
of oxidase activity (Fig. 4). None of the fragments recovered from the EcoRI or PstI
digestions of p1S11D were active.
Although the DNA inserts in p1U12G and p1S11D are not identical, their
restriction enzyme digestion patterns suggested that they share a
common ~9-kb segment of DNA (data not shown). In fact, the insert in
p1U12G contains a ~6.5-kb HindIII fragment (Fig. 3) that
appears to be identical to the one from p1S11D, as judged by the fact
that it also can direct the expression of oxidase activity (data not
shown). Thus, p1U12G and p1S11D appear to share overlapping DNA
segments containing the oxidase structural gene.
Identification of orfE (lpxQ) as the Structural Gene for the
Oxidase--
Based upon DNA sequencing, at least nine complete or
partial open reading frames were tentatively identified on the 6.5-kb HindIII fragment present in pQN210 (Fig.
5). Sequence similarity searches with the
BLASTx program indicated that orfA and orfM (Fig.
5) encode glycolate oxidase subunits. However, when orfA was
cloned into pRK404a and then transferred into S. meliloti, no lipid A oxidase activity was observed in cell extracts (data not
shown).
Both OrfC and OrfD (Fig. 5) share significant sequence similarity with
a set of hypothetical membrane proteins of unknown function found in
various members of the Rhizobiacea and other bacteria, including
R. leguminosarum, Mesorhizobium loti, and S. meliloti. Expression cloning of orfC and
orfD (either separately or together) in pRK404a and
subsequent transfer into S. meliloti failed to induce
oxidase activity in cell extracts (data not shown). A similar hybrid
plasmid harboring orfB, which encodes a protein with strong
similarity to DNA-3-methyladenine glycosylase I, was likewise inactive
in S. meliloti.
Although OrfE does not show strong similarity to any functionally
assigned protein in the NCBI data base, it does display weak similarity
to an outer surface protein of Wolbachia and to the outer
membrane ferripyoverdine receptor of Pseudomonas. As discussed further below, a significant OrfE ortholog of unknown function is present in the plant pathogen A. tumefaciens
(Fig. 6). Interestingly, extracts of
S. meliloti/pQN231, which contains R. leguminosarum
orfE behind a lac promoter on pRK404a, display robust
oxidase activity (Fig. 7). Additional
constructs containing orfE in various T7lac
promoter-driven pET vectors (designated pQN233 through pQN235 in Table
I) direct high levels of oxidase expression in E. coli cell
extracts and membranes (Fig. 7), providing unequivocal evidence that
orfE is the oxidase structural gene. Given its unique
function in lipid A modification, we suggest that orfE be
renamed lpxQ in accordance with the nomenclature used for
other genes encoding lipid A biosynthetic enzymes (16, 38, 39).
Expression and Function of the lpxQ Ortholog of A. tumefaciens in
E. coli--
As shown in Fig. 6, the genomes of both sequenced strains
of A. tumefaciens encode a protein that is ~59% identical
and ~77% similar to R. leguminosarum LpxQ (17, 18).
Expression of the putative A. tumefaciens lpxQ gene behind
the T7lac promoter in two strains of E. coli
resulted in the appearance of significant lipid A oxidase activity in
cell extracts, as judged by the conversion of [14C]B to
[14C]D-1 (Fig.
8). This finding suggests that A. tumefaciens may be able to synthesize 2-aminogluconate containing
lipid A under some conditions.
Mass Spectrometry of D-1 Synthesized in Vitro by Recombinant
LpxQ--
The membranes of E. coli that express R. leguminosarum LpxQ were used to convert component B to a substance
resembling D-1, as judged by TLC. The latter product was then separated
from residual B and purified by DEAE cellulose column chromatography,
as described in the accompanying article (1) for D-1 synthesized by
membranes of R. leguminosarum 3855. MALDI/TOF mass
spectrometry of the product in the positive mode (Fig.
9B) reveals ions at
m/z 1996.2 and 2024.6, which are interpreted as
[M+Na]+ of D-1 species differing in acyl chain length.
Fig. 9A shows the positive mode spectrum of an authentic
sample of D-1 isolated from R. etli CE3 (3), which gives
rise to [M + Na]+ ions at m/z
1996.2 and 2024.6, both of which are 16 atomic mass units larger than
the corresponding species in component B (spectrum not shown). The D-1
standard and the material synthesized by LpxQ under these conditions
both possess identical B Disappearance of LpxQ Activity under Anaerobic Conditions in
Vitro--
As discussed in the accompanying article (1), a plausible
mechanism for the oxidation B to D-1 might involve oxygen
as the electron accepting substrate. Although it has not yet been possible to demonstrate stoichiometric hydrogen peroxide formation concomitant with D-1 synthesis, the results of Fig.
10 clearly demonstrate that LpxQ
activity is strictly dependent upon the presence of oxygen. The further
addition of glucose oxidase and catalase did not alter the rate of
D-1 formation. Even after an overnight incubation, no
detectable product was observed during oxygen deprivation (data not
shown). The apparent requirement of recombinant LpxQ for molecular
oxygen provides the strongest evidence presently available that this
enzyme might be a novel type of oxidase.
LpxQ Localizes to the Outer Membrane when Expressed in E. coli--
Isopycnic sucrose density gradient centrifugation of
membranes prepared from induced cells of Novablue(DE3)/pQN233 was used to evaluate the subcellular localization of the recombinant LpxQ oxidase. The inner and outer membranes of the induced construct were
well separated, as shown in Fig.
11A, by assay of the marker enzymes NADH oxidase and phospholipase A, respectively. Most of the
lipid A oxidase is associated with the outer membrane in this construct
(Fig. 11B), as in R. leguminosarum 3855 (1).
A protein corresponding in size to that expected for LpxQ (~23 kDa)
is present in the outer membranes of Novablue(DE3)/pQN233 (Fig.
12), as judged by SDS-PAGE, but not in
the outer membranes of the vector control Novablue(DE3)/pET21a+. The
program SignalP (40) does in fact predict that LpxQ is an outer
membrane protein with an N-terminal signal peptide that may be cleaved
between Ala27 and Glu28. Signal
sequences of this kind are present in virtually all outer membrane
proteins of Gram-negative bacteria, because they are essential for
proper translocation (41). The N-terminal sequence of the first 10 amino acids of the putative mature LpxQ protein band present in the
outer membranes of Novablue(DE3)/pQN233 was determined as
EDLQFSIYGG. This result corresponds precisely to the predicted cleavage
site and establishes conclusively that LpxQ is a genuine outer
membrane protein.
All Gram-negative bacteria synthesize the lipid A
component of lipopolysaccharide by a means of a constitutive seven-step pathway that starts with UDP-GlcNAc and proceeds via the tetra-acylated precursor, Kdo2-lipid IVA, as indicated
schematically in Fig. 13 (16, 42). At
least one additional secondary acyl chain is usually added after Kdo
incorporation, most commonly at the 2' position (16, 42). In the case
of R. leguminosarum, this additional acyl chain is 28 carbon
atoms long and is hydroxylated at position 27 (Fig. 1) (1, 43, 44). A
special acyl carrier protein is required for 27-OH-C28 synthesis
and transfer to lipid A (Fig. 13) (43, 44). In E. coli a
secondary laurate chain is added at the 2' position, and myristate is
added at 3' (1, 45-48). All of the reactions leading to
Kdo2-lipid IVA are cytosolic or associated with
the inner membrane, as is the incorporation of 27-OH-C28 (Fig. 13) in
R. leguminosarum or laurate and myristate in E. coli (16, 42).
INTRODUCTION
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
Bacterial strains and plasmids used in this study
80 °C as master stocks in the same dishes
after adjusting the medium to 20% glycerol by addition of an
appropriate volume of 60% glycerol. To prepare fresh lysates for
screening purposes, 1800 colonies were taken out of storage and regrown
in 96-well microtiter plates. Each well was inoculated with a 5-µl
portion of the master glycerol stock culture into 150 µl of TY broth
supplemented with 10 mM CaCl2, 20 µg/ml
nalidixic acid, 200 µg/ml streptomycin sulfate, and 12 µg/ml
tetracycline. The microtiter plates were then shaken at 225 rpm in a
30 °C incubator for ~24 h or until the OD550
reached ~ 0.6, as measured with a microtiter plate reader.
(Back-up glycerol stocks of these master plates were made by
transferring 50 µl from each well into a new microtiter plate, in
which each well contained 25 µl of 60% glycerol. The new stocks were
stored at
80 °C.)
80 °C freezer for several hours, after which they were thawed.
Each 96-well microtiter plate contained eight rows (rows A-H) of 12 wells (wells 1-12). To facilitate the screening, each row was further
grouped into four pools of three lysates. Row A, for example, was
grouped into pools A(1-3), A(4-6), A(7-9), and A(10-12). The pooled
lysates were assayed for their ability to convert [14C]B
to [14C]D-1. A portion of each pool (9 µl) was mixed
with 2 µl of a concentrated reaction buffer (consisting of 5 mM MgCl2, 250 mM MES, pH 6.5, and
0.5% Triton X-100) and 0.5 µl of 0.05 µM
[14C]B (~600 cpm/reaction tube). After 30 and 60 min of
incubation at 30 °C, 5-µl portions of each reaction mixture were
spotted onto a 20 × 20-cm silica TLC plate. Negative (no enzyme)
and positive (wild type R. leguminosarum 3855 membranes)
controls were included for each 96-well microtiter plate of pooled
lysates that were screened. The amount of [14C]D-1 formed
in each reaction tube was quantified with a PhosphorImager.
80 °C prior to
preparation of membranes. Two other E. coli host strains, BL21(DE3)/pLysS and Novablue(DE3), were tested in the same manner for
their ability to express the lpxQ gene product. The empty vector pET21b+ was also transformed into each of the above host strains
in parallel.
RESULTS
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ABSTRACT
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EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 1.
Expression cloning of the R. leguminosarum lipid A oxidase in R. etli. A, a genomic R. leguminosarum DNA library in cosmid pLAFR-1 (24, 32, 44) was
transferred into R. etli CE3. Cells from ~1800
cosmid-containing colonies were grown up individually in 96-well
microtiter plates. As described under "Experimental Procedures,"
the lysates prepared by lysozyme treatment were pooled into groups of
three and assayed for overexpression of the lipid A oxidase activity,
as judged by the conversion of ~0.003 µM
[14C]B (~600 cpm/reaction tube) to
[14C]D-1 after 60 min. The arrow indicates a
possible pool with elevated oxidase activity. B, the
calculated percentage of conversion of [14C]B to
[14C]D-1 is shown for each lane. The activity in the pool
from row D (wells 10-12 from plate 1S containing lysates 1S10D, 1S11D,
and 1S12D) is approximately twice that of the other pools. Assays of
the individual lysates (data not shown) revealed that only 1S11D
contained high levels of oxidase activity. Among the ~1800 colonies
tested, only three positive cosmids (p1S11D, p1E11D, and p1U12G) were
identified in this manner. Based on restriction enzyme digests, p1S11D
and p1E11D contained the same insert.
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Fig. 2.
Transfer of the R. leguminosarum
cosmids expressing the lipid A oxidase activity into S. meliloti. The active cosmids identified in Fig. 1 and
the empty vector pLAFR-1 were transferred into S. meliloti
1021 via tri-parental mating (27, 28). Cell-free extracts and membranes
derived from late log phase cells containing these cosmids were
prepared and assayed for oxidase activity using 10 µM
[14C]B (600 cpm/reaction tube) as the substrate.
Lanes 2-4 show the results with 0.5 mg/ml crude extracts,
whereas lanes 5-7 show assays with 0.5 mg/ml washed
membranes. Lane 1, no enzyme control; lane 2,
pLAFR-1; lane 3, p1S11D; lane 4, p1U12G;
lane 5, pLAFR-1; lane 6, p1S11D; lane
7, p1U12G. The reaction mixtures were incubated overnight at
30 °C. Similar results were obtained when
[4'-32P]1-dephospho-lipid IVA was used as the
substrate (data not shown).
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Fig. 3.
Digestion of cosmids p1S11D, p1U12G, and
pRK404a with EcoRI and HindIII.
Cosmids p1U12G (lanes 2-4) and p1S11D (lanes
7-9) were digested with EcoRI (E),
HindIII (H), or with both EcoRI and
HindIII (E/H). The vector pRK404a was digested in
parallel (lanes 5 and 6). The HindIII
fragment migrating near 6.5 kb (indicated by the thin line)
directed the overexpression of oxidase activity (see below).
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Fig. 4.
A 6.5-kb HindIII digestion
fragment of p1S11D directs the overexpression of the oxidase. The
cosmid p1S11D harbors ~20 kb of R. leguminosarum 8401 genomic DNA. Subclones of the insert were constructed by restriction
enzyme digestion and ligation of the fragments into pRK404a. Plasmids
pQN209-pQN214 were derived from various HindIII fragments
of the p1S11D insert. pQN208 contains a 3.9-kb EcoRI
fragment of the insert, and pQN215 contains a ~0.5-kb PstI
fragment. All of the constructs were transferred into S. meliloti 1021 by tri-parental mating (27, 28). Only pQN210, which
contains the ~6.5-kb HindIII fragment of the p1S11D insert
(see Fig. 3) directed the overexpression of oxidase activity in
S. meliloti, as shown by assaying 0.5 mg/ml washed membranes
from cells harboring the various constructs with 5 µM
[14C]B for 30 min at 30 °C.
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Fig. 5.
Order of the R. leguminosarum
genes present on the DNA insert in plasmid pQN210. Open
reading frames contained within the 6.5-kb insert present in pQN210
were identified based on analysis of the DNA sequence (accession number
AY228164) with the program ORF Finder (31) and compared with the
nonredundant data base with BLASTx (54). The most plausible candidate
for the oxidase is the lpxQ gene. H,
HindIII; B, BamHI; Sm,
SmaI; Sl, SalI; P,
PstI; E, EcoRI.
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Fig. 6.
Sequence comparison of LpxQ from R. leguminosarum and Agrobacterium
tumefaciens. The predicted amino acid sequence of the
R. leguminosarum lipid A oxidase LpxQ (accession number
AY228164) is compared with an ortholog of unknown function from
A. tumefaciens (17, 18). The order and sequence of the other
genes around LpxQ is likewise conserved in both organisms. No other
proteins with significant similarity to LpxQ are present in the NCBI
data base.
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Fig. 7.
Heterologous expression of the
lpxQ encoded oxidase in S. meliloti
and E. coli. The lpxQ gene was
amplified by PCR and ligated into both the shuttle vector pRK404a and
into the T7-promoter based vector pET21a+. The resulting constructs,
pQN231 and pQN233, were transferred into S. meliloti 1021 and E. coli BL21(DE3)/pLysS, respectively. S. meliloti/pQN231 membranes were prepared from late log phase cells.
Membranes of BL21(DE3)/pLysS/pQN233 were obtained from mid log phase
cells induced with 1 mM IPTG for 3 h. The membranes
(0.5 mg/ml) were assayed for 120 min at 30 °C for their ability to
convert 10 µM [14C]B to
[14C]D-1 under standard oxidase assay conditions.
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Fig. 8.
The lpxQ homolog of A. tumefaciens directs the expression of oxidase activity in
E. coli. The lpxQ homolog of A. tumefaciens was amplified by PCR and cloned into pET21b+. The
resulting hybrid plasmid is designated pQN240. Membranes of E. coli BL21(DE3)/pLysS/pQN240 (lanes 2 and 3)
or BLR(DE3)/pLysS/pQN240 (lanes 5 and 6), grown
and induced as described in the legend to Fig. 7, were assayed under
standard conditions for 15 min. The no enzyme control is shown in
lane 1. Membranes derived from cells containing the vector
were assayed in parallel (lanes 4 and 7). A
positive control (i.e. membranes of the R. leguminosarum lpxQ overexpressing strain E. coli
BL21(DE3)/pLysS/pQN233) is shown in lane 8.
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Fig. 9.
Positive ion MALDI/TOF mass spectrum of D-1
generated by membranes of BL21(DE3)/pLysS/pQN233. A reaction
mixture (5 ml) containing 50 µM B was incubated for
16 h at 30 °C with 0.5 mg/ml membranes of
BL21(DE3)/pLysS/pQN233 under standard conditions, as described in the
accompanying article (1). Following partial purification of the
D-1-like reaction product by ion exchange chromatography on a small
column of DEAE-cellulose, MALDI/TOF mass spectrometry was performed in
the positive ion mode (1). Panel A shows the spectrum of a
standard preparation of component D-1 isolated from R. etli
(3). Panel B shows the spectrum of the partially purified
in vitro reaction product. In addition to confirming the
incorporation of an oxygen atom into the proximal unit of B, this
spectrum also reveals that about half of the D-1 was further modified
with a palmitate residue, presumably because of the PagP
acyltransferase activity that is present in membranes of the E. coli host strain (7).
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Fig. 10.
The lipid A oxidase encoded by
lpxQ is dependent upon atmospheric oxygen. The
oxidase assay was performed either in an anaerobic chamber or under
ambient atmospheric conditions, using membranes of an E. coli strain expressing a C-terminal His-tagged version of
LpxQ. The conversion of [14C]B to [14C]D-1
was determined after 30, 60, or 90 min at 30 °C, using 0.1 mg/ml
membranes from strain BLR(DE3)/pLysS/pQN235 that was induced with
IPTG as in Fig. 5. The presence of glucose oxidase and catalase had
little or no effect on the rate of the reaction or the extent of
conversion.
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Fig. 11.
Outer membrane localization of LpxQ oxidase
activity in E. coli Novablue(DE3)/pQN233. Washed
membranes obtained from induced cells of E. coli
Novablue(DE3)/pQN233 were separated by isopycnic sucrose density
gradient centrifugation. Marker enzymes localized in the outer or inner
membranes (phospholipase A and NADH oxidase respectively) were assayed
to evaluate the extent of separation. A, phospholipase A and
NADH oxidase activity. B, LpxQ oxidase activity and protein
concentration.
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Fig. 12.
Outer membrane localization of overexpressed
LpxQ protein in E. coli
Novablue(DE3)/pQN233. Membranes of E. coli
Novablue(DE3)/pQN233 and Novablue(DE3)/pET21a+ were fractionated by
isopycnic sucrose gradient centrifugation as described under
"Experimental Procedures." Peak fractions of outer and inner
membranes were pooled separately, and recovered by ultracentrifugation.
The membranes were subjected to SDS-PAGE (12% gel and 40 µg of
protein/lane). A band of the size expected for LpxQ is seen only in the
outer membranes derived from E. coli Novablue(DE3)/pQN233,
as indicated by the arrow. Lane 1, Molecular weight marker
(Benchmark Protein Ladder from Invitrogen); lane 2, outer
membranes of Novablue(DE3)/pQN233; lane 3, outer
membranes of Novablue(DE3)/pET21a+; lane 4, inner membranes
of Novablue(DE3)/pQN233; lane 5, inner membranes of
Novablue(DE3)/pET21a+. The putative LpxQ band, indicated by the
arrow, was excised and subjected to N-terminal
microsequencing.
DISCUSSION
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Fig. 13.
Proposed compartmentalization of R. leguminosarum lipid A biosynthesis and function of
LpxQ. Almost all of the enzymes needed to generate component B
have been detected in extracts of R. leguminosarum and
R. etli (43, 44, 55-57). The only exceptions are the
reactions that incorporate the 4'-galacturonic acid and the
-hydroxybutyrate residues. Although the hypothetical glycolipid,
undecaprenyl-phosphate-GalUA, has not actually been isolated from cells
or confirmed as the GalUA donor substrate in vitro, recent
work in our laboratory suggests that GalUA transfer to R. leguminosarum lipopolysaccharide precursors requires a
membrane-bound donor (S. S. Basu, M. Kanipes, and C. R. H. Raetz, unpublished results). The lpxE gene, which was
recently found by expression cloning,2 encodes the
1-phosphatase, and it is predicted to have a periplasmic active site.
The gene encoding the 4'-phosphatase is unknown. The ABC transporter
MsbA is proposed to catalyze the flip-flop of the nascent lipid A 1,4'
bis-phosphate with attached core sugars across the inner
membrane (58-60), thereby presenting this intermediate to the
periplasmic lipid A modification enzymes. Following the formation of
component B and transport to the outer membrane by unknown mechanisms,
the LpxQ oxidase converts the proximal glucosamine residue to the
2-aminogluconate unit in an oxygen-dependent manner. If the
oxidation reaction proceeds through a lactone intermediate (as
discussed in the accompanying article (1)), an additional lactonase
(not shown) might be needed to generate D-1. However, hydrolysis of
such a lactone could be catalyzed by LpxQ itself or be
nonenzymatic.
An emerging theme in the enzymatic assembly of lipid A is that additional (often regulated) covalent modifications of the conventional lipid A disaccharide 1, 4'-bis-phosphate backbone may occur beyond the cytoplasm (16). In polymyxin-resistant mutants of E. coli and S. typhimurium, the 4-amino-4-deoxy-L-arabinose (L-Ara4N) substituent is attached to the 4' phosphate moiety of lipid A on the outer surface of the inner membrane (49, 50). The proposal is supported by the recent discovery that the L-Ara4N donor substrate is the novel lipid, undecaprenyl phosphate-L-Ara4N, and by the predicted topography of the enzyme (ArnT) that transfers the L-Ara4N unit to lipid A (49, 50). Some covalent modifications of lipid A can even occur in the outer membrane. The addition of a secondary palmitate chain at the 2 position of lipid A is catalyzed by the outer membrane acyltransferase PagP in S. typhimurium and E. coli, which uses a phospholipid as its acyl donor (7, 8, 51). The recent NMR studies of PagP have localized the active site of the enzyme to the outer surface of the outer membrane (8). Finally, S. typhimurium also contains PagL, a specific lipid A 3-O deacylase that is localized in the outer membrane (34). Unlike the constitutive (cytoplasmic) enzymes in the lipid A pathway (Fig. 13), which is highly conserved in virtually all Gram-negative bacteria (16), the lipid A-modifying enzymes found in the outer layers of the cell envelope are variable and relatively restricted in their distribution.
The data presented in this and the accompanying article (1) demonstrate conclusively that LpxQ is a novel type of outer membrane enzyme catalyzing an unusual oxidative modification of lipid A molecules that are dephosphorylated at the 1 position (Fig. 13). Heterologous expression behind the T7lac promoter in E. coli provides definitive evidence that lpxQ is indeed the structural gene for the oxidase (Fig. 7). Although two other outer membrane enzymes (PagP and PagL) have recently been discovered that catalyze covalent modifications of lipid A in S. typhimurium and E. coli (7, 34), as noted above, LpxQ is the first example of an outer membrane oxidase in any bacterial system.
The primary sequence of LpxQ is distinct from all previously described sugar oxidases or dehydrogenases but is distantly similar to selected outer membrane proteins, as judge by BLASTp or PSI-BLAST analysis of the nonredundant data base. The only significant ortholog (Fig. 6) of LpxQ (55% identity and 77% similarity over the length of the protein) currently in the nonredundant data base occurs in strains of A. tumefaciens, the plant pathogen that causes Crown-Gall disease and that is used to generate transgenic and mutant plants (17, 18). The results of Fig. 8 show that A. tumefaciens LpxQ expressed in E. coli catalyzes the same oxidative lipid A modification as does R. leguminosarum LpxQ. However, the presence of lipid A species containing the 2-aminogluconate moiety have not actually been described in A. tumefaciens, although this issue has not been investigated in any depth. Interestingly, the identity and arrangement of the genes that flank LpxQ in A. tumefaciens (17, 18) is quite similar to what is seen in R. leguminosarum (Fig. 5).
The function of the 2-aminogluconate unit generated by LpxQ in R. leguminosarum is unknown. The occurrence of the lpxQ gene in two important bacterial systems that are of great interest in plant biology necessitates further studies of the significance of the 2-aminogluconate moiety. The isolation of mutants of R. leguminosarum and A. tumefaciens, in which the lpxQ gene is deleted, should provide a powerful approach to this problem, especially if such strains are viable and grow normally under laboratory conditions. These mutants could be tested for their ability to infect and interact with their plant hosts in comparison with wild type bacteria. The additional availability of the lpxE gene encoding the lipid A 1-phosphatase,2 which generates the substrate for LpxQ (Fig. 13), together with lpxQ might even enable the oxidative modification of lipid A in bacteria like S. meliloti, which do not ordinarily make the 2-aminogluconate residue. The R. leguminosarum lpxE structural gene encoding the 1-phosphatase has recently been cloned in our laboratory, and its properties will be described elsewhere.2 It is an inner membrane protein with an active site that is predicted to face the periplasm (Fig. 13).
Our results demonstrate that formation of the 2-aminogluconate unit in component D-1 occurs late in the R. leguminosarum lipid A pathway. The lipid A of R. leguminosarum/pQN210, which overexpresses lpxQ, is highly enriched in component D-1 with very little remaining B (data not shown). In contrast, the ratio between B to D-1 in wild type R. leguminosarum lipid A is usually about 1:2 (3, 4). The extra copies of the oxidase present in R. leguminosarum/pQN210 appear to deplete the available component B.
The availability of the lpxQ gene should facilitate the
purification of the oxidase to homogeneity. With pure enzyme it should be possible to explore the proposed role of oxygen as the electron accepting cosubstrate in the conversion of B to D-1 and to confirm the
stoichiometric formation of hydrogen peroxide. The identification of
possible organic cofactors, the characterization of the catalytic mechanism, and the evaluation of the significance of the EDTA inhibition should be greatly simplified with pure protein. Finally, the
structural biology of LpxQ should also be of great interest in relation
to other outer enzymes of know structure, like PagP (8), OmpT (9), and
PldA (6). The tertiary structure of LpxQ might reveal how the
characteristic inside out -barrel folds of outer membrane proteins
have evolved to generate novel catalytic sites.
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ACKNOWLEDGEMENTS |
---|
We thank Kimberly Johnson and Margo Wuebbens of the Rajagopalan laboratory at Duke University for assistance with the anaerobic chamber.
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FOOTNOTES |
---|
* This work was supported by National Institutes of Health Grants R37-GM-51796 (to C. R. H. R.) and GM54882 (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.
§ Supported by National Institutes of Health Training Grant GM-08558 in Biological Chemistry to Duke University.
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.M300379200
2 M. Karbarz and C. R. H. Raetz, manuscript in preparation.
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ABBREVIATIONS |
---|
The abbreviations used are:
MES, 2-(N-morpholino)-ethanesulfonic acid;
2-aminogluconate, 2-amino-2-deoxy-gluconate;
CAPS, 3-(cyclohexylamino)propanesulfonic
acid;
IPTG, isopropyl-1-thio--D-galactopyranoside;
MALDI, matrix-assisted laser desorption ionization;
TOF, time-of-flight;
Kdo, 3-deoxy-D-manno-2-octulosonic acid.
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REFERENCES |
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![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. |
Que-Gewirth, N. L. S.,
Lin, S.,
Cotter, R. J.,
and Raetz, C. R. H.
(2003)
J. Biol. Chem.
278,
12109-12119 |
2. |
Bhat, U. R.,
Forsberg, L. S.,
and Carlson, R. W.
(1994)
J. Biol. Chem.
269,
14402-14410 |
3. |
Que, N. L. S.,
Lin, S.,
Cotter, R. J.,
and Raetz, C. R. H.
(2000)
J. Biol. Chem.
275,
28006-28016 |
4. |
Que, N. L. S.,
Ribeiro, A. A.,
and Raetz, C. R. H.
(2000)
J. Biol. Chem.
275,
28017-28027 |
5. | Nishijima, M., Nakaike, S., Tamori, Y., and Nojima, S. (1977) Eur. J. Biochem. 73, 115-124[Abstract] |
6. | Snijder, H. J., Ubarretxena-Belandia, I., Blaauw, M., Kalk, K. H., Verheij, H. M., Egmond, M. R., Dekker, N., and Dijkstra, B. W. (1999) Nature 401, 717-721[Medline] [Order article via Infotrieve] |
7. |
Bishop, R. E.,
Gibbons, H. S.,
Guina, T.,
Trent, M. S.,
Miller, S. I.,
and Raetz, C. R. H.
(2000)
EMBO J.
19,
5071-5080 |
8. |
Hwang, P. M.,
Choy, W. Y.,
Lo, E. I.,
Chen, L.,
Forman-Kay, J. D.,
Raetz, C. R. H.,
Prive, G. G.,
Bishop, R. E.,
and Kay, L. E.
(2002)
Proc. Natl. Acad. Sci. U. S. A.
99,
13560-13565 |
9. |
Vandeputte-Rutten, L.,
Kramer, R. A.,
Kroon, J.,
Dekker, N.,
Egmond, M. R.,
and Gros, P.
(2001)
EMBO J.
20,
5033-5039 |
10. |
Ferguson, A. D.,
Hofmann, E.,
Coulton, J. W.,
Diederichs, K.,
and Welte, W.
(1998)
Science
282,
2215-2220 |
11. | Koronakis, V., Sharff, A., Koronakis, E., Luisi, B., and Hughes, C. (2000) Nature 405, 914-919[CrossRef][Medline] [Order article via Infotrieve] |
12. | Schulz, G. E. (2002) Biochim. Biophys. Acta 1565, 308-317[Medline] [Order article via Infotrieve] |
13. | Fernandez, C., Hilty, C., Bonjour, S., Adeishvili, K., Pervushin, K., and Wüthrich, K. (2001) FEBS Lett. 504, 173-178[CrossRef][Medline] [Order article via Infotrieve] |
14. |
Groisman, E. A.
(2001)
J. Bacteriol.
183,
1835-1842 |
15. | Ohl, M. E., and Miller, S. I. (2001) Annu. Rev. Med. 52, 259-274[CrossRef][Medline] [Order article via Infotrieve] |
16. | Raetz, C. R. H., and Whitfield, C. (2002) Annu. Rev. Biochem. 71, 635-700[CrossRef][Medline] [Order article via Infotrieve] |
17. |
Wood, D. W.,
Setubal, J. C.,
Kaul, R.,
Monks, D. E.,
Kitajima, J. P.,
Okura, V. K.,
Zhou, Y.,
Chen, L.,
Wood, G. E.,
Almeida, N. F., Jr.,
Woo, L.,
Chen, Y.,
Paulsen, I. T.,
Eisen, J. A.,
Karp, P. D.,
Bovee, D., Sr.,
Chapman, P.,
Clendenning, J.,
Deatherage, G.,
Gillet, W.,
Grant, C.,
Kutyavin, T.,
Levy, R.,
Li, M. J.,
McClelland, E.,
Palmieri, A.,
Raymond, C.,
Rouse, G.,
Saenphimmachak, C.,
Wu, Z.,
Romero, P.,
Gordon, D.,
Zhang, S.,
Yoo, H.,
Tao, Y.,
Biddle, P.,
Jung, M.,
Krespan, W.,
Perry, M.,
Gordon-Kamm, B.,
Liao, L.,
Kim, S.,
Hendrick, C.,
Zhao, Z. Y.,
Dolan, M.,
Chumley, F.,
Tingey, S. V.,
Tomb, J. F.,
Gordon, M. P.,
Olson, M. V.,
and Nester, E. W.
(2001)
Science
294,
2317-2323 |
18. |
Goodner, B.,
Hinkle, G.,
Gattung, S.,
Miller, N.,
Blanchard, M.,
Qurollo, B.,
Goldman, B. S.,
Cao, Y.,
Askenazi, M.,
Halling, C.,
Mullin, L.,
Houmiel, K.,
Gordon, J.,
Vaudin, M.,
Iartchouk, O.,
Epp, A.,
Liu, F.,
Wollam, C.,
Allinger, M.,
Doughty, D.,
Scott, C.,
Lappas, C.,
Markelz, B.,
Flanagan, C.,
Crowell, C.,
Gurson, J.,
Lomo, C.,
Sear, C.,
Strub, G.,
Cielo, C.,
and Slater, S.
(2001)
Science
294,
2323-2328 |
19. | Cava, J. R., Elias, P. M., Turowski, D. A., and Noel, K. D. (1989) J. Bacteriol. 171, 8-15[Medline] [Order article via Infotrieve] |
20. | Segovia, L., Young, J. P., and Martinez-Romero, E. (1993) Int. J. Syst. Bacteriol. 43, 374-377[Abstract] |
21. | Meade, H. M., Long, S. R., Ruvkun, G. B., Brown, S. E., and Ausubel, F. M. (1982) J. Bacteriol. 149, 114-122[Medline] [Order article via Infotrieve] |
22. |
Galibert, F.,
Finan, T. M.,
Long, S. R.,
Puhler, A.,
Abola, P.,
Ampe, F.,
Barloy-Hubler, F.,
Barnett, M. J.,
Becker, A.,
Boistard, P.,
Bothe, G.,
Boutry, M.,
Bowser, L.,
Buhrmester, J.,
Cadieu, E.,
Capela, D.,
Chain, P.,
Cowie, A.,
Davis, R. W.,
Dreano, S.,
Federspiel, N. A.,
Fisher, R. F.,
Gloux, S.,
Godrie, T.,
Goffeau, A.,
Golding, B.,
Gouzy, J.,
Gurjal, M.,
Hernandez-Lucas, I.,
Hong, A.,
Huizar, L.,
Hyman, R. W.,
Jones, T.,
Kahn, D.,
Kahn, M. L.,
Kalman, S.,
Keating, D. H.,
Kiss, E.,
Komp, C.,
Lelaure, V.,
Masuy, D.,
Palm, C.,
Peck, M. C.,
Pohl, T. M.,
Portetelle, D.,
Purnelle, B.,
Ramsperger, U.,
Surzycki, R.,
Thebault, P.,
Vandenbol, M.,
Vorholter, F. J.,
Weidner, S.,
Wells, D. H.,
Wong, K.,
Yeh, K. C.,
and Batut, J.
(2001)
Science
293,
668-672 |
23. | Miller, J. R. (1972) Experiments in Molecular Genetics , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY |
24. | Ronson, C. W., Astwood, P. M., and Downie, J. A. (1984) J. Bacteriol. 160, 903-909[Medline] [Order article via Infotrieve] |
25. | Friedman, A. M., Long, S. R., Brown, S. E., Buikema, W. J., and Ausubel, F. M. (1982) Gene (Amst.) 18, 289-296[CrossRef][Medline] [Order article via Infotrieve] |
26. | Wood, W. (1966) J. Mol. Biol. 16, 118-133[Medline] [Order article via Infotrieve] |
27. | Glazebrook, J., and Walker, G. C. (1991) Methods Enzymol. 204, 398-418[Medline] [Order article via Infotrieve] |
28. | Finan, T. M., Kunkel, B., De Vos, G. F., and Signer, E. R. (1986) J. Bacteriol. 167, 66-72[Medline] [Order article via Infotrieve] |
29. | Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual , 2nd Ed. , Cold Spring Harbor Laboratory, Cold Spring Harbor, NY |
30. | Bergmans, H. E., van Die, I. M., and Hoekstra, W. P. (1981) J. Bacteriol. 146, 564-570[Medline] [Order article via Infotrieve] |
31. |
Wheeler, D. L.,
Church, D. M.,
Federhen, S.,
Lash, A. E.,
Madden, T. L.,
Pontius, J. U.,
Schuler, G. D.,
Schriml, L. M.,
Sequeira, E.,
Tatusova, T. A.,
and Wagner, L.
(2003)
Nucleic Acids Res.
31,
28-33 |
32. | Ditta, G., Stanfield, S., Corbin, D., and Helinski, D. R. (1980) Proc. Natl. Acad. Sci. U. S. A. 77, 7347-7351[Abstract] |
33. | MacFerrin, K. D., Terranova, M. P., Schreiber, S. L., and Verdine, G. L. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 1937-1941[Abstract] |
34. |
Trent, M. S.,
Pabich, W.,
Raetz, C. R. H.,
and Miller, S. I.
(2001)
J. Biol. Chem.
276,
9083-9092 |
35. | Guy-Caffey, J. K., Rapoza, M. P., Jolley, K. A., and Webster, R. E. (1992) J. Bacteriol. 174, 2460-2465[Abstract] |
36. | Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985) Anal. Biochem. 150, 76-85[Medline] [Order article via Infotrieve] |
37. | Laemmli, U. K. (1970) Nature 227, 680-685[Medline] [Order article via Infotrieve] |
38. | Raetz, C. R. H. (1990) Annu. Rev. Biochem. 59, 129-170[CrossRef][Medline] [Order article via Infotrieve] |
39. | Raetz, C. R. H. (1993) J. Bacteriol. 175, 5745-5753[Medline] [Order article via Infotrieve] |
40. |
Nielsen, H.,
Brunak, S.,
and von Heijne, G.
(1999)
Protein Engineering
12,
3-9 |
41. | Wickner, W., Driessen, A. J., and Hartl, F. U. (1991) Annu. Rev. Biochem. 60, 101-124[CrossRef][Medline] [Order article via Infotrieve] |
42. | Raetz, C. R. H. (1996) in Escherichia coli and Salmonella: Cellular and Molecular Biology (Neidhardt, F. C., ed), 2nd Ed., Vol. 1 , pp. 1035-1063, American Society for Microbiology, Washington, D.C. |
43. |
Brozek, K. A.,
Carlson, R. W.,
and Raetz, C. R. H.
(1996)
J. Biol. Chem.
271,
32126-32136 |
44. |
Basu, S. S.,
Karbarz, M. J.,
and Raetz, C. R. H.
(2002)
J. Biol. Chem.
277,
28959-28971 |
45. |
Clementz, T.,
Bednarski, J. J.,
and Raetz, C. R. H.
(1996)
J. Biol. Chem.
271,
12095-12102 |
46. |
Clementz, T.,
Zhou, Z.,
and Raetz, C. R. H.
(1997)
J. Biol. Chem.
272,
10353-10360 |
47. |
Vorachek-Warren, M. K.,
Carty, S. M.,
Lin, S.,
Cotter, R. J.,
and Raetz, C. R. H.
(2002)
J. Biol. Chem.
277,
14186-14193 |
48. |
Vorachek-Warren, M. K.,
Ramirez, S.,
Cotter, R. J.,
and Raetz, C. R. H.
(2002)
J. Biol. Chem.
277,
14194-14205 |
49. |
Trent, M. S.,
Ribeiro, A. A.,
Lin, S.,
Cotter, R. J.,
and Raetz, C. R. H.
(2001)
J. Biol. Chem.
276,
43122-43131 |
50. |
Trent, M. S.,
Ribeiro, A. A.,
Doerrler, W. T.,
Lin, S.,
Cotter, R. J.,
and Raetz, C. R. H.
(2001)
J. Biol. Chem.
276,
43132-43144 |
51. |
Brozek, K. A.,
Bulawa, C. E.,
and Raetz, C. R. H.
(1987)
J. Biol. Chem.
262,
5170-5179 |
52. | Ronson, C. W., Astwood, P. M., Nixon, B. T., and Ausubel, F. M. (1987) Nucleic Acids Res. 15, 7921-7934[Abstract] |
53. | Russa, R., Lüderitz, O., and Rietschel, E. T. (1985) Arch. Microbiol. 141, 284-289 |
54. | Gish, W., and States, D. J. (1993) Nat. Genet. 3, 266-272[Medline] [Order article via Infotrieve] |
55. | Price, N. P. J., Kelly, T. M., Raetz, C. R. H., and Carlson, R. W. (1994) J. Bacteriol. 176, 4646-4655[Abstract] |
56. | Price, N. J. P., Jeyaretnam, B., Carlson, R. W., Kadrmas, J. L., Raetz, C. R. H., and Brozek, K. A. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 7352-7356[Abstract] |
57. |
Brozek, K. A.,
Kadrmas, J. L.,
and Raetz, C. R. H.
(1996)
J. Biol. Chem.
271,
32112-32118 |
58. |
Zhou, Z.,
White, K. A.,
Polissi, A.,
Georgopoulos, C.,
and Raetz, C. R. H.
(1998)
J. Biol. Chem.
273,
12466-12475 |
59. |
Doerrler, W. T.,
Reedy, M. C.,
and Raetz, C. R. H.
(2001)
J. Biol. Chem.
276,
11461-11464 |
60. |
Chang, G.,
and Roth, C. B.
(2001)
Science
293,
1793-1800 |