JOURNAL OF BACTERIOLOGY, Jan. 2007, p. 244–253
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
Vol. 189, No. 1
Characterization of lptA and lptB, Two Essential Genes Implicated
in Lipopolysaccharide Transport to the Outer Membrane
of Escherichia coli?
Paola Sperandeo,1Rachele Cescutti,2Riccardo Villa,2Cristiano Di Benedetto,3Daniela Candia,3
Gianni Deho `,1and Alessandra Polissi2*
Dipartimento di Scienze Biomolecolari e Biotecnologie, Universita ` degli Studi di Milano, Milan, Italy1; Dipartimento di
Biotecnologie e Bioscienze, Universita ` di Milano-Bicocca, Milan, Italy2; and Dipartimento di Biologia,
Universita ` degli Studi di Milano, Milan, Italy3
Received 27 July 2006/Accepted 11 October 2006
The outer membrane (OM) of gram-negative bacteria is an asymmetric lipid bilayer that protects the cell
from toxic molecules. Lipopolysaccharide (LPS) is an essential component of the OM in most gram-negative
bacteria, and its structure and biosynthesis are well known. Nevertheless, the mechanisms of transport and
assembly of this molecule in the OM are poorly understood. To date, the only proteins implicated in LPS
transport are MsbA, responsible for LPS flipping across the inner membrane, and the Imp/RlpB complex,
involved in LPS targeting to the OM. Here, we present evidence that two Escherichia coli essential genes, yhbN
and yhbG, now renamed lptA and lptB, respectively, participate in LPS biogenesis. We show that mutants
depleted of LptA and/or LptB not only produce an anomalous LPS form, but also are defective in LPS transport
to the OM and accumulate de novo-synthesized LPS in a novel membrane fraction of intermediate density
between the inner membrane (IM) and the OM. In addition, we show that LptA is located in the periplasm and
that expression of the lptA-lptB operon is controlled by the extracytoplasmic ? factor RpoE. Based on these
data, we propose that LptA and LptB are implicated in the transport of LPS from the IM to the OM of E. coli.
The cell envelope of gram-negative bacteria consists of an
inner (IM) and an outer (OM) membrane separated by the
periplasmic space, which contains the peptidoglycan layer. The
OM is an asymmetric bilayer, with phospholipids in the inner
leaflet and lipopolysaccharides (LPS) facing outward (28, 32).
LPS, a molecule unique to gram-negative bacteria, consists of
the lipid A moiety (a glucosamine-based phospholipid) linked
to a short-core oligosaccharide and the distal O-antigen poly-
saccharide chain. The core oligosaccharide can be further
divided into the inner core, composed of 3-deoxy-D-manno-
octulosanate (KDO) and heptose, and the outer core, with a
somewhat variable structure. Escherichia coli K-12 synthe-
sizes a shorter LPS consisting of a lipid A moiety linked to
a short-core oligosaccharide but missing the O-antigen
chain (28). LPS is essential in most gram-negative bacteria,
with the notable exception of Neisseria meningitidis. In E.
coli, the minimal part required for viability consists of the
KDO2lipid IVAmoiety (28).
LPS is synthesized in the cytoplasmic face of the IM and
must traverse the IM and the periplasm to reach its final
destination in the outer face of the OM (6, 28). Translocation
across the IM requires the ABC transporter MsbA, which
mediates the flipping from the inner leaflet (the site of the
synthesis) to the outer leaflet of the IM (14, 27, 44). MsbA has
also been implicated in phospholipid transport across the IM
of E. coli (13, 14). Interestingly, MsbA in N. meningitidis is not
essential and seems not to be required for phospholipid trans-
port (39). How LPS traverses the periplasm and is inserted into
the OM is much less understood.
Because of its hydrophobic lipid A moiety, transport of LPS
through the aqueous periplasm is thermodynamically unfavor-
able. In E. coli, it has been shown that lipoprotein (Lpp)
transport across the periplasmic space is mediated by the LolA
chaperone protein. The system is energized by the LolCDE
protein complex, an ABC transporter located in the IM. After
transport across the periplasm, Lpps are transferred from
LolA to the receptor, LolB, in the OM (reviewed in reference
41). No evidence for a similar mechanism has been found so
far in the transport of LPS. de Cock and coworkers have
recently shown in E. coli spheroplasts that de novo-synthesized
LPS, unlike Lpps, are not released in the presence of a
periplasmic extract. Moreover, the authors showed that LPS
cofractionated with remnants of the OM, and this was depen-
dent on the presence of a functional MsbA protein in sphero-
plasts. These observations have been taken as evidence that
transport of LPS could proceed via contact sites between the
two membranes (40), thus lending support to earlier proposals
that IM-OM adhesion sites (also known as Bayer bridges)
could be implicated in LPS transport to the OM (references 2
and 3 and references therein; 16). However, the existence of
these points of physical contact between the two membranes
has been questioned; they may be artifacts occurring during
the cell fixation process used in electron microscopy (17).
A protein implicated in LPS targeting to the OM is Imp, a
?-barrel OM protein. Braun and Silhavy demonstrated that
Imp is essential in E. coli and that its depletion results in
abnormalities in OM assembly, with newly synthesized lipids
and outer membrane proteins appearing in a novel membrane
* Corresponding author. Mailing address: Dipartimento di Biotec-
nologie e Bioscienze, Universita ` di Milano-Bicocca, Piazza della Sci-
enza 220126, Milano, Italy. Phone: 39-02-64483431. Fax: 39-02-6448-
3450. E-mail: email@example.com.
?Published ahead of print on 20 October 2006.
fraction with higher density than the OM (7). Bos and cowork-
ers demonstrated that Imp is required for proper transport of
LPS to the cell surface of N. meningitidis (5). This conclusion
was based upon the loss of surface accessibility by LPS to
neuraminidase and loss of lipid A modification by the OM
deacylase PagL (5). However, as in this organism it is not
possible to separate the IM and OM by sucrose gradient ultra-
centrifugation, it could not be assessed where LPS accumu-
lates. In agreement with the observation that N. meningitidis
can survive without LPS, imp is not an essential gene in this
organism (5). Very recently, the essential E. coli lipoprotein
RlpB has been shown to interact physically with Imp. RlpB
depletion results in defects in OM biogenesis similar to those
of Imp depletion, and based on the observation that depletion
of both proteins activates the OM enzyme PagP to modify LPS,
the authors conclude that the Imp/RlpB complex is responsible
for LPS reaching the outer surface of the OM (43).
The OM represents an effective permeability barrier to pro-
tect the cells from toxic compounds, such as antibiotics and
detergents, thus allowing bacteria to inhabit several different
and often hostile environments. The integrity of the extracy-
toplasmic compartments (the periplasm and OM) is monitored
by at least two signaling systems, dependent on Cpx and ?E,
respectively, that sense the status of the cell envelope and
respond to repair damage when necessary (reviewed in ref-
erence 33). The Cpx system seems to be triggered mainly by
damage to surface structures, such as pili (29), whereas the
alternative ?Efactor is activated by unfolded envelope pro-
teins and abnormal LPS (20, 23, 38). In agreement with this,
genes implicated in cell envelope biogenesis (such as imp) or
in LPS biosynthesis have been found to belong to the ?E
regulon (7, 10).
We have recently identified two new essential genes of E.
coli, yhbN and yhbG, with unknown functions (35). yhbN and
yhbG are the distal genes of a cluster comprising four addi-
tional genes. Several lines of evidence obtained in our labora-
tory implicate these two genes in OM biogenesis: (i) yhbN and
yhbG are both essential and cotranscribed, (ii) they are well
conserved in many gram-negative bacteria, (iii) the cluster in
which they are organized comprises two genes (kdsD and
kdsC) involved in KDO biosynthesis, and (iv) mutants defec-
tive in their expression exhibit altered OM permeability (35).
In this study, we provide evidence that YhbN and YhbG are
implicated in the transport of LPS from the inner to the outer
membrane of E. coli. Henceforth, yhbN and yhbG will be re-
named lptA and lptB (for LPS transport), respectively.
MATERIALS AND METHODS
Bacterial strains and media. The bacterial strains and plasmids used in this
work are listed in Table 1. Oligonucleotide primers are listed in Table 2. Bacteria
were grown in either LD broth or M9 minimal medium (18) supplemented with
0.2% glycerol as a carbon source. When required, 0.2% L-arabinose (as an
inducer of the araBp promoter), 0.2% glucose, 20 ?g/ml Casamino Acids, 0.5
mM IPTG (isopropyl-?-D-thiogalactopyranoside), 100 ?g/ml ampicillin (Amp),
50 ?g/ml kanamycin (Kan), and 25 ?g/ml chloramphenicol (Cam) were added.
Solid media were as described above with 1% agar.
Construction of mutants BB-10 and BB-11 by allele replacement. Allele re-
placement was performed using the procedure based on the ? Red recombina-
tion system (11). To obtain BB-10 (Table 1), the mutant ?rseA1::cat allele was
prepared in vitro by a three-step PCR procedure (8) using AP97 and AP98
primers and, as templates, the cat cassette (which confers chloramphenicol re-
sistance) amplified by pKD3 using AP79-AP80 primers and two DNA fragments
for the flanking homology regions obtained by PCR amplification of E. coli
MG1655 DNA with oligonucleotides AP95-AP97 and AP96-AP98, respectively
(Table 2). Insertion of the cat cassette resulted in the removal of amino acid
residues 2 to 198 of the 216-codon-long rseA coding sequence. To obtain the
?rseA1 mutant (BB-11), BB-10 was transformed with pCP20 and ampicillin-
resistant transformants were selected at 30°C. Removal of the cat cassette, which
generated an in-frame deletion replaced by an 18-codon scar (11), was then
obtained by colony purification of a few transformants at 43°C and screening
them for loss of both Amp and Cam resistance. The presence of the ?rseA1 allele
in BB-11 was verified by PCR.
Plasmid construction. Plasmids pGS109 and pGS110 express LptA and LptB,
respectively, with a C-terminal His6tag (LptA-H and LptB-H) upon IPTG
induction. They were constructed by PCR amplifying the lptA and lptB open
reading frames from genomic MG1655 DNA with primers AP55-AP64 (lptA) or
AP56-AP65 (lptB) (Table 2). The PCR products were EcoRI-XbaI digested and
cloned in pGS100 cut with the same enzymes. The EcoRI-XbaI inserts in the
pGS109 and pGS110 were verified by sequencing.
Plasmids pRP1 and pRP2 were constructed by cloning into EcoRI-BamHI-
digested pRS415 the E. coli chromosomal regions containing the putative lptAp
and yrbGp promoters, respectively. The lptAp and yrbGp DNAs were PCR
amplified from MG1655 genomic DNA using primers AP69-AP70 and AP71-
AP72, respectively, and digested with EcoRI-BamHI. The two EcoRI-BamHI
inserts in plasmids pRP1 and pRP2 were verified by sequencing.
Total LPS extraction and analysis. Bacterial cultures grown at 37°C in LD-
arabinose up to an optical density at 600 nm (OD600) of 0.2 were harvested by
centrifugation, washed in LD, and diluted 100-fold in LD with or without arabi-
nose. Samples at a total OD600of 2 were taken at different time points, and LPS
was extracted from cell pellets by a mini-phenol-water extraction technique as
described previously (30). Briefly, the cells were resuspended in water and
pelleted (5 min; 10,000 ? g) to remove the exopolysaccharides. The pellet was
TABLE 1. Bacterial strains and plasmids
BW25113 ?(kan araC araBp-yhbN)1
BW25113 ?(kan araC araBp-yhbG)1
BW25113 ?rseA1; in-frame deletion
by removal of cat cassette from
?(argF-lac169) ?80 dlacZ58(M15)
glnV44(AS) ??rfbD1 gyrA96
recA1 endA1 spoT1 thi-1 hsdR17
pGS100pGZ119EH derivative; contains TIR
sequence downstream of tacp
FLP ?cI857 ?P Rep(Ts) CamrAmpr
oriR?; AmprCamr; source of cat
oriR101 repA101(Ts) araC araBp-
pBR322-derived plasmid containing
the complete lac operon without
pRS415 derivative containing the
?E-dependent lptAp promoter
pRS415 derivative containing the
?70-dependent yrbGp promoter
VOL. 189, 2007CHARACTERIZATION OF E. COLI lptA AND lptB245
resuspended in 0.3 ml phosphate buffer, pH 7, and thoroughly vortexed; 0.3 ml
of phenol equilibrated with 0.1 M Tris-HCl at pH 5.5 was added, and the
suspension was vortexed. The tubes were placed in a 65°C heating block for 15
min with thorough vortexing every 5 min and then cooled in ice. After centrif-
ugation (5 min; 10,000 ? g), the water phase was removed, dialyzed (2,000-
molecular-weight cutoff) against phosphate buffer, pH 7, and dried under vac-
uum. The lyophilized material was then dissolved in 30 ?l of polyacrylamide gel
electrophoresis (PAGE) sample buffer. LPS was separated by Tricine–sodium
dodecyl sulfate (SDS)-PAGE and silver stained as described previously (36).
N-Acetyl[3H]glucosamine pulse-labeling and cell fractionation. Mutants were
grown in LD-arabinose up to an OD600of 0.2 at 37°C in the presence of 0.2%
N-acetylglucosamine (GlnNAc) to induce GlnNAc uptake. The wild-type strain
was grown under the same conditions, except that arabinose was omitted from
the medium. Cells were then harvested, washed in LD, diluted 100-fold in fresh
medium (50 ml) with or without arabinose, and incubated with aeration at 37°C.
[3H]GlnNAc pulse-labeling (see below) of the wild type and mutants grown with
arabinose was performed when the cultures reached an OD600of 0.6, whereas
depleted cultures grown without arabinose were labeled 1 h after the cells
reached their maximal OD600(between 0.4 and 0.6). [3H]GlnNAc pulse-labeling
was performed by adding N-acetyl[3H]glucosamine (1.6 ?Ci ml?1) to the culture
at 37°C and diluting the culture with 1 volume of the same medium containing
0.8% nonradioactive GlnNAc after 2 min. After a 5-min chase, the cells were
chilled in ice and harvested by centrifugation. The inner and outer membranes
were separated by discontinuous sucrose density gradient centrifugation of a
total membrane fraction obtained by spheroplast lysis as described previously
(26). Step gradients were prepared by layering 2 ml each of 50, 45, 40, 35, and
30% sucrose solutions (wt/vol) over a 55% sucrose cushion (0.5 ml).
Fractions (200 ?l) were collected from the top of the gradient, and 50 ?l each
was assayed for NADH oxidase activity (26). The total protein concentration was
determined in 10-?l samples of each fraction by the Bradford assay (Bio-Rad) as
recommended by the manufacturer. The protein profiles of OmpC, OmpF, and
OmpA across the gradient were estimated by separating 20 to 40 ?l of each
fraction on 12.5% SDS-PAGE and by staining the gels with Coomassie blue. To
estimate3H incorporation, 25 ?l of each fraction dissolved in 5 ml scintillation
liquid (Ready Safe; Beckman-Coulter) was counted in a liquid scintillation
counter (LS6500; Beckman).
Analysis of LPS from gradient fractions. To estimate the LPS contents in the
gradient fractions, equal volumes were digested with 6 ?g of proteinase K
(Sigma) at 60°C for 1 h and then separated by the Tricine–SDS-PAGE procedure
as described previously (36). LPS were transferred onto a nitrocellulose mem-
brane (Hybond ECL; GE Healthcare) or a polyvinylidene difluoride micro-
porous membrane (Immobilon-P; Millipore) at 60 V for 2 h in an electroblotting
apparatus (PBI). LPS on nitrocellulose membranes were immunodetected using
a 1:10,000 dilution of the anti-LPS WN1 222-5 monoclonal antibody (HyCult
Biotechnology b.v.). Polyvinylidene difluoride membranes with radiolabeled LPS
were scanned with a Beta Imager 2000 (Biospace).
Cellular localization of LptA. Cultures of BW25113 containing plasmid
pGS109 were grown in M9 glucose minimal medium up to an OD600of 0.7, and
LptA-H was induced for 1 h with 0.5 mM IPTG. Periplasmic, cytoplasmic, inner,
and outer membrane fractions were prepared as described previously (25). Equal
amounts of proteins from each fraction were fractionated by 12.5% SDS-PAGE.
The tagged proteins were detected by Western blotting using monoclonal anti-
?-Galactosidase assay. Overnight cultures of bacterial strains harboring pRP1,
pRP2, or pRS415 plasmids in LD at 30°C were diluted 1:100 and grown with
aeration up to an OD600of 0.4. The ?-galactosidase activity was then determined
by the method of Miller (22).
Light microscopy. BB-4 cells grown in LD-arabinose up to an OD600of 0.2
were harvested by centrifugation, washed in LD, and diluted 100-fold in LD with
or without arabinose. Growth was monitored by reading the OD600, and samples
were taken at different time points. Cells were harvested by centrifugation,
washed, and resuspended in phosphate-buffered saline. The cells were imaged
with a Leica TCS SP2 confocal microscope coupled to a Leica DMIRE2 inverted
microscope. Transmission images were obtained by the Ar laser line at 488 nm
through a PL APO 63? oil immersion objective (numerical aperture, 1.4).
Electron microscopy. Bacterial samples obtained as described above were
pelleted in Eppendorf tubes, washed with cacodylate buffer (0.2 M, pH 7.4), and
fixed with 2% glutaraldehyde in 0.1 M cacodylate buffer. Samples were then
postfixed with 1% osmium tetroxide in 0.1 M cacodylate buffer, dehydrated in a
graded ethanol series, and embedded in an Epon-Araldite mixture according
to standard transmission electron microscopy (TEM) methods (19). Semithin
(1-?m) and ultrathin (?50-nm) sections were cut with a Reichert-Jung
ULTRACUT E using diamond knives (DIATOME histo and Ultra 45°). Semi-
thin sections, collected on standard slides and stained with crystal violet and basic
fuchsin, were controlled under a Jenaval light microscope. Ultrathin sections,
collected on 300-mesh copper grids, were stained with aqueous uranyl acetate
and lead citrate (31), carbon coated under an EMITECH K400X carbon coater,
and observed with a Jeol 100 SX electron microscope. Micrographs were taken
directly under the microscope with Kodak 4489 photographic films for TEM.
Expression of lptA and lptB is controlled by ?E. Genetic
analysis of the yrbG-lptB locus indicated that lptA and lptB are
organized in a dicistronic operon, and a putative ?E-dependent
TABLE 2. Oligonucleotides
Name5? to 3? sequencea
pGS109 construction, with AP64; EcoRI
pGS110 construction, with AP65; EcoRI
pGS109 construction, with AP55; insertion
of C-terminal His6tag into LptA; XbaI
pGS110 construction, with AP56; insertion
of C-terminal His6tag into LptB; XbaI
pRP1 construction, with AP70; EcoRI
pRP1 construction, with AP69; BamHI
pRP2 construction, with AP72; EcoRI
pRP2 construction, with AP71; BamHI
Amplification of cat cassette from pKD3 with AP80
Amplification of cat cassette from pKD3 with AP79
For control of BB-10 and BB-11 mutant regions,
For control of BB-10 and BB-11 mutant regions,
cat hybrid primer for BB-10 construction by
three-step PCR, with AP97
cat hybrid primer for BB-10 construction by
three-step PCR, with AP98
BB-10 construction by three-step PCR, with AP95
BB-10 construction by three-step PCR, with AP96
aE. coli genomic sequences are in uppercase; restriction sites are underlined and specified in the “Use/description” column.
246SPERANDEO ET AL.J. BACTERIOL.
promoter consensus sequence was identified 61 nucleotides
upstream of the lptA start codon (35) (Fig. 1). To obtain ex-
perimental evidence of this putative promoter, we created in
the promoter-probe vector pRS415 an lptAp-lacZ and, as a
control, a yrbGp-lacZ translational fusion (pRP1 and pRP2,
respectively) (Table 1). The ?-galactosidase activity was as-
sessed in BW25113 and its ?rseA1 derivative, BB-11 (Table 1).
RseA is an anti-? factor whose loss results in a constitutively
high level of ?Eactivity (24). The ?-galactosidase activity in
BB-11 transformed with pRP1 was approximately 10-fold
higher than the activity measured in BW25113 transformed
with the same plasmid, whereas no variation was observed in
both BW25113 and BB-11 harboring either pRP2 or the pro-
moterless vector (Fig. 2). Thus, lptA and lptB can be expressed
from a ?E-dependent promoter.
LptA is a periplasmic protein. Recently, LptB (YhbG) has
been found associated with the plasma membrane (37),
whereas nothing was known about the cellular localization of
LptA. The lptA gene encodes a 185-amino acid protein (pre-
dicted mass, 20.13 kDa). Primary sequence analysis revealed a
putative N-terminal signal sequence, suggesting that the pro-
tein is transported across the IM. Blast homology searches
found LptA homologues in all five subclasses of Proteobacteria
but none in gram-positive bacteria and in Archea (http://www
.ncbi.nlm.nih.gov/BLAST/). None of these putative proteins
has been characterized. To define the subcellular localization
of LptA, we constructed a tagged version of the protein car-
rying a C-terminal His6tag (LptA-H) under the control of the
IPTG-inducible tacp promoter (plasmid pGS109) (Table 1).
Ectopic expression of the LptA protein in rich medium was
toxic for cells (not shown), and thus, growth and induction with
IPTG were performed in M9-glucose minimal medium.
Periplasmic, cytoplasmic, inner, and outer membrane fractions
from IPTG-induced and noninduced BW25113/pGS109 were
prepared as described previously (25) and analyzed by Western
blotting using monoclonal anti-His6tag antibodies. As shown
in Fig. 3, LptA-H was detectable only in the periplasmic frac-
Morphological and structural abnormalities in LptA-LptB-
depleted cells. BB-4 is a conditional-lethal (arabinose-depen-
dent) mutant in which the arabinose-inducible araBp promoter
drives expression of lptA and lptB (Table 1). LptA-LptB de-
pletion leads to growth arrest and cell death after six or seven
generations (150 min following the shift to a medium lacking
arabinose), followed by a modest decline in cell turbidity (35)
(Fig. 4A). We thus examined whether LptA-LptB depletion
would cause recognizable morphological anomalies in mutant
cells. Light microscopy observation showed no difference be-
tween depleted and nondepleted cells up to 90 min after the
removal of arabinose. At later time points (150 and 210 min),
LptA-LptB-depleted cells showed mostly short filaments (Fig.
4B). Furthermore, ultrastructural analysis by TEM allowed us
to detect accumulated extra membrane material or multilay-
ered membranous bodies within the periplasmic space and the
budding of vesicles derived from the OM (Fig. 4D and E). A
similar phenotype has been observed in Imp- and RlpB-de-
pleted cells (43).
We then examined the LPS profile. Total LPS was extracted
at 90, 150, and 210 min; fractionated by Tricine–SDS-PAGE;
and silver stained. Interestingly, in depleted BB-4 cells late
(150 and 210 min) after arabinose removal, an anomalous form
of LPS visible as a ladder of bands migrating more slowly than
the native LPS was observed (Fig. 4C). In addition, increased
levels of LPS seemed to accumulate in the BB-4 mutant 210
min after LptA-LptB depletion (Fig. 4C), as was also observed
upon Imp and RlpB depletion (43).
Depletion of LptA and LptB affects LPS transport to the
OM. To further investigate the roles of LptA and LptB in LPS
maturation, we followed the fate of newly synthesized LPS by
FIG. 1. Organization of the yrbG-lptB locus. The open reading frames are drawn to scale as arrowed rectangles (open, nonessential genes; solid,
essential genes; gray, truncated flanking genes). Promoters are indicated by bent arrows (open, putative promoters; solid, documented promoters).
Coordinates (in kb) on the top bar are from the E. coli complete genomic sequence (GenBank NC_000913) after subtracting 3337.
FIG. 2. Promoter activities of yrbGp and lptAp. The activities of the
reporter fusions yrbGp-lacZ and lptAp-lacZ were assessed in wild-type
and ?rseA strains. ?-Galactosidase activity measurements were per-
formed in duplicate and calculated in Miller units. The data presented
are the averages (with standard deviations) of three independent ex-
FIG. 3. Subcellular localization of LptA. BW25113 cells containing
plasmid pGS109 were induced with IPTG and then fractionated as
described in Materials and Methods. P, periplasmic proteins; S, cyto-
plasmic proteins; IM, solubilized inner membrane proteins; OM, sol-
ubilized outer membrane proteins; T, total proteins.
VOL. 189, 2007 CHARACTERIZATION OF E. COLI lptA AND lptB 247
pulse-labeling wild-type, BB-4-depleted, and nondepleted cells
with the LPS precursor N-acetyl[3H]glucosamine. Membranes
were then fractionated by isopycnic sucrose density gradient
centrifugation, as described in Materials and Methods. The
fractions were assayed for total proteins, NADH oxidase ac-
tivity as an IM marker, incorporated radioactive precursor
(Fig. 5A, C, and E), total LPS,3H-labeled LPS, and OmpC,
OmpF, and OmpA porin profiles (Fig. 5B, D, and F). As
shown in Fig. 5A and C, wild-type and nondepleted BB-4 cells
displayed a bimodal protein distribution. Most NADH oxidase
activity was found in the lighter peak (IM; fractions 15 to 23),
whereas the denser peak (OM; fractions 39 to 43) contained
most of the total LPS (Fig. 5B and D, LPS) and proteins of
43.3, 39.1, and 37 kDa, corresponding to the masses of OmpC,
OmpF, and OmpA porins, respectively, as seen by SDS-PAGE
(Fig. 5B and D, OmpC/F-OmpA). The radioactivity in wild-
type and nondepleted BB-4 cells was found mainly associated
with fractions corresponding to the OM, indicating that
N-acetyl[3H]glucosamine was readily incorporated into LPS
(Fig. 5A and C). Moreover,
fractions 39 to 45, indicating that newly synthesized LPS are
transported to the OM (Fig. 5B and D,3H-LPS).
3H-labeled LPS were found in
On the other hand, the fractionation profiles of LptA-LptB-
depleted cells were markedly different. Proteins did not display
a clear bimodal pattern, and a peak of intermediate density
containing mostly NADH oxidase activity appeared (Fig. 5E,
heavy IM [hIM], fractions 29 to 33). The major amount of LPS
was found in fractions denser than the hIM (fractions 33 to 43).
However, the distribution of LPS was skewed toward lower
densities than those found in wild-type and nondepleted cells.
These data seem to indicate that a substantial fraction of the
IM and OM cannot be cleanly separated by buoyant-density
centrifugation. This is further supported by the distribution of
OmpC/F-OmpA proteins, which were detected by SDS-PAGE
in fractions 33 to 41, thus floating at a density lower than the
OM (Fig. 5F, OmpC/F-OmpA).
Interestingly, a substantial amount of the abnormal form of
high-molecular-weight LPS could be detected in fractions 15 to
27 and was thus lighter than hIM. In the denser fractions,
however, the presence of the abnormal LPS could be masked
by the strong LPS signal. Moreover the distribution of radio-
activity was found mainly associated with the hIM peak of
intermediate density (Fig. 5E, fractions 29 to 33). Finally, as
expected from the radioactivity profile, most of de novo-syn-
FIG. 4. Growth curve, cell morphology, and LPS profile upon LptA-LptB depletion. (A) Bacterial cultures grown in LD-arabinose up to an
OD600of 0.2 were harvested by centrifugation, washed in LD, and diluted 100-fold in LD with arabinose (?ara) or without arabinose (?ara).
Growth was monitored by optical density at 600 nm. (B) Light microscopy images of LptA-LptB-depleted (?ara) and nondepleted (?ara) cells
taken at different time points. (C) LPS profiles of BB-4 and BW25113 (wild type [WT]) grown with and without arabinose. LPS extracted from
cultures with a total OD600of 2 were separated on 18% Tricine–SDS-PAGE and silver stained. The amount of LPS corresponding to an OD600
of 0.4 was loaded in each lane. (D and E) Electron microscopy images of wild-type (D) and BB-4 depleted (E) cells. Scale bars, 0.5 ?m.
248 SPERANDEO ET AL. J. BACTERIOL.
thesized LPS fractionated in the lighter part of the gradient
corresponding to IM and hIM peaks, whereas little label was
detected in heavier fractions (Fig. 5F,3H-LPS). These data
indicate that LptA-LptB depletion results in a dramatic alter-
ation of both the IM and the OM and that under these con-
ditions, newly synthesized LPS are not transported to the OM.
Both LptA and LptB are implicated in LPS transport to
the OM. The experiments described above show that repres-
sion of the lptA-lptB operon impairs LPS transport and OM
biogenesis. We thus tested whether each gene is implicated
in this process by analyzing the LPS-incorporated radioac-
tivity profile in an N-acetyl[3H]glucosamine pulse-chase/
membrane fractionation experiment, as described above, in
cultures individually depleted of either LptA or LptB. LptB
depletion was achieved in BB-5 (araBp-lptB), in which the
expression of lptB is arabinose dependent (35), whereas for
depletion of LptA, we complemented BB-4 (araBp-lptA-
lptB) with pGS110, a plasmid that expresses LptB-H (i.e.,
FIG. 5. Membrane fractionation and LPS profile of LptA-LptB-depleted mutant. BB-4 mutant cells shifted to an arabinose-free medium were
pulse-labeled for 2 min with [3H]GlnNAc 1 hour after they had reached their maximal OD600(about 0.5 to 0.6) and then harvested by
centrifugation 5 min after the chase; the nondepleted and wild-type cultures were pulse-labeled when they reached the same OD600, as described
in Materials and Methods. Total membranes prepared from cells were fractionated into inner and outer membranes by sucrose density gradient
centrifugation. Fractions of 200 ?l were collected from the top of the gradient and analyzed for total protein content, NADH oxidase activity,
OmpC/F-OmpA protein profiles, and total and labeled LPS. (A, C, and E) Membrane fractionation profiles of BW25113, nondepleted BB-4, and
LptA-LptB-depleted BB-4 cells, respectively. NADH oxidase activity (?) and disintegration per minutes (DPM; black diamonds) are expressed
as percentages of the total. Total proteins (open circles), determined with the Bio-Rad Bradford assay, are expressed in mg/ml. (B, D, and F) LPS
profiles of BW25113, nondepleted BB-4, and LptA-LptB-depleted BB-4 cells, respectively. LPS, LPS profiles of fractions determined by Western
blotting using anti-LPS WN1 222-5 monoclonal antibody;3H-LPS, newly synthesized radiolabeled LPS scanned with a ?-imager; OmpC/F-OmpA,
profiles of the major OM porins.
VOL. 189, 2007 CHARACTERIZATION OF E. COLI lptA AND lptB249
with a C-terminal His6tag) (Table 1). pGS110 was able to
complement the arabinose dependence of BB-5, even in the
absence of IPTG (data not shown), indicating that the LptB-H
is functional and sufficiently expressed even in the absence if
When membranes were separated from nondepleted BB-5
and BB-4/pGS110, the fractionation profiles were very sim-
ilar to those seen in BW25113 or in nondepleted BB-4 cells.
As judged by the NADH oxidase activity profile, the IM
sedimented around fractions 11 to 19 in both mutants (Fig.
6A and C, top), whereas, based on both total-protein and
porin profiles, OM equilibrated around fractions 39 to 43
(Fig. 6A and C, top and bottom). The distribution of radio-
activity in BB-5 and BB-4/pGS110 nondepleted cells was
found mainly associated with the fractions corresponding to
the OM (Fig. 6A and C, top), indicating that the newly
synthesized LPS is transported to the OM. On the other
hand, when either LptA or LptB was depleted, the fraction-
ation profile resembled that observed in BB-4 depleted cells.
Most of the NADH oxidase activity appeared in a peak of
intermediate density (hIM fractions 21 to 31 and 21 to 33 in
BB4/pGS110 and BB-5, respectively). Although the total-
protein profile did not display a clear bimodal pattern, a
small peak around fractions 39 to 41 with a density corre-
sponding to the OM and containing porins appeared in both
BB4/pGS110 and BB-5 (Fig. 6B and D, top). However, in
both depleted mutants, the distribution of porins was
skewed toward lower densities than those found in nonde-
pleted cells (Fig. 6B and D, bottom). Interestingly, the ra-
dioactivity was found to be associated mainly with the peak
of intermediate density in both mutants (fractions 21 to 31
and 21 to 33 in BB4/pGS110 and BB-5, respectively), and
very little radioactivity was found in the fractions corre-
sponding to the OM, thus indicating that most of the de
novo-synthesized LPS are not transported to the OM.
Overall, these data indicate that depletion of either lptA or
lptB is sufficient to cause a dramatic alteration of both the IM
and OM with concomitant impairment of LPS targeting to the
OM, and thus, both genes seem to act in the same pathway of
LPS transport across the periplasmic space.
FIG. 6. Membrane fractionation of LptA- or LptB-depleted cells. BB-4/pGS110 and BB-5 mutant cells shifted to an arabinose-free medium
were pulse-labeled for 2 min with [3H]GlnNAc 1 hour after they had reached their maximal OD600(about 0.4 to 0.6) and then harvested by
centrifugation 5 min after the chase; the nondepleted cultures were pulse-labeled when they reached the same OD600, as described in Materials
and Methods. Total membranes prepared from cells were fractionated into inner and outer membranes by sucrose density gradient centrifugation.
Fractions of 200 ?l were collected from the top of the gradient and analyzed for total protein content, NADH oxidase activity, radioactivity profiles,
and OmpC/F-OmpA protein profiles. (A and B) Membrane fractionation profiles and the profiles of the major OM porins (OmpC/F-OmpA) of
nondepleted and depleted BB-5 cells, respectively. (C and D) Membrane fractionation profiles and the profiles of the major OM porins
(OmpC/F-OmpA) of nondepleted and depleted BB-4/pGS110 cells, respectively. NADH oxidase activity (?) and disintegrations per minutes
(DPM; black diamonds) are expressed as percentages of the total. Total proteins (open circles), determined with the Bio-Rad Bradford assay, are
expressed in mg/ml.
250 SPERANDEO ET AL.J. BACTERIOL.
In this work, we implicate the E. coli essential genes lptA and
lptB in LPS transport across the periplasm. We show that the
two genes are expressed from a ?E-dependent promoter. The
?Eregulon responds to envelope stress and is specifically ac-
tivated by both misfolded OM proteins and alteration of LPS
structure (33, 38). Among the target genes whose expression is
upregulated by ?E, there are factors implicated in OM biogen-
esis. These include Imp, with a postulated role in LPS assembly
in the OM (7); SurA, a periplasmic chaperone necessary for
proper OM protein folding (7, 10); and YaeT, YfgL, YfiO, and
NlpB, which form a multiprotein complex required for OM
protein assembly (32). In the yrbG-lptB locus, two additional
promoters, kdsCp and the ?70-dependent yrbGp, are predicted
upstream of lptAp (35; this work) (Fig. 1), and we cannot rule
out the possibility that the lptA and lptB genes may also be
expressed from these upstream promoters. However, the pres-
ence of a ?E-dependent promoter places lptA and lptB in the
growing family of genes with extracytoplasmic functions.
We demonstrated that upon LptA-LptB depletion, [3H]Glc-
NAc, a compound that is normally incorporated into the LPS
outer core, accumulates in a novel membrane fraction with
higher density than the IM (hIM) and is not transported to the
OM. Moreover, in depleted cells,3H-LPS fractionates with the
hIM fraction and is absent in denser fractions (Fig. 5). Similar
defects are observed in cells in which LptA and LptB are
individually depleted (Fig. 6), indicating that both proteins are
required for LPS targeting to the OM. Electron micrographs of
LptA-LptB-depleted cells also show serious envelope defects
resembling those observed upon Imp or RlpB depletion (43).
These data suggest that LptA and LptB are OM biogenesis
factors implicated in LPS transport.
We do not know the nature of the hIM peak appearing upon
LptA-LptB depletion. A likely hypothesis is that LPS accumu-
lates in the IM, increasing the LPS/phospholipid ratio and thus
increasing the density of the IM fraction, whereas conversely,
the blockade of LPS transport across the periplasm may result
in decreased density of the OM. The behavior of LptA-LptB-
depleted cells is reminiscent of Imp and RlpB depletion, where
newly synthesized lipids and proteins accumulated in a novel
membrane fraction with increased density over that of the OM,
which has been taken as evidence of an increased LPS/phos-
pholipid ratio (7, 43). Alternatively, the blockade of the LPS
transport machinery may cause abnormal membrane aggrega-
tions, as was also observed by electron microscopy, that pre-
vent a clean separation of the IM and OM through sucrose
LPS accumulation in LptA-LptB- and Imp-RlpB-depleted
cells suggests that the rate of LPS transport is not coupled to
the rate of its synthesis. Interestingly a similar observation was
made by Meredith and coworkers, who found that in a mutant
unable to synthesize KDO, thus producing only lipid IVA, the
rate of lipid IVAtransport had become uncoupled from its rate
of synthesis (21). However, LPS accumulation and changes in
membrane fractionation profiles have not been observed upon
MsbA depletion or thermal inactivation of a temperature-sen-
sitive msbA allele (13, 27, 44). It is possible, however, that this
difference could be imputed to different experimental condi-
tions (e.g., incubation under nonpermissive conditions was not
long enough for this to occur).
The pulse-labeling experiments with LptA-LptB-depleted
cells using [3H]GlcNAc were unable to distinguish the topol-
ogy of LPS in the hIM peak. However, based on the published
data on MsbA-dependent LPS translocation, we hypothesize
that in the LptA-LptB-depleted mutants, LPS accumulates in
the outer leaflet of the IM. This is consistent with the electron
microscopy observation of extra membrane accumulated in the
periplasmic space, as was also observed in RlpB- and Imp-
depleted mutants (43).
LptA-LptB-depleted cells produce an abnormal, higher-mo-
lecular-weight form of LPS that can be detected both in total
extracts and in fractionated membranes. Following transloca-
tion across the IM, the KDO2lipid IVAmoiety with the core
sugars attached may undergo additional chemical modifica-
tions either in the periplasm or in the OM (4, 42). In the
periplasm, the covalent modification of lipid A with the cat-
ionic sugar 4-amino-4-deoxy-L-arabinose catalyzed by ArnT is
induced by environmental stimuli or appropriate point muta-
tions and leads to polymyxin resistance (42). The anomalous
LPS produced upon LptA-LptB depletion could be the result
of such chemical modifications occurring when the molecule
stacks in the IM facing the periplasm. Further analysis will test
LptA, as suggested by its amino acid sequence analysis, is a
periplasmic protein (Fig. 3) predicted to work with LptB, a
protein belonging to the superfamily of ABC transporters
(http://ecocyc.org/). Amino acid sequence analysis suggests
that the 26.6-kDa LptB protein is soluble and possesses the
ATP binding fold, but not the transmembrane domain. How-
ever, LptB has been recently identified in the IM in a com-
plex of ?140 kDa, although no interacting partners could be
detected (37). We suggest (see the model in Fig. 7) that
LptA and LptB, together with an as-yet-unidentified trans-
membrane partner, may form a membrane-associated com-
plex required for LPS transport, possibly by providing the
required energy. A transmembrane partner candidate is
YrbK, an essential protein encoded by the gene immediately
FIG. 7. A model for the transport of LPS. LPS (dark ovals with six
small wavy lines, lipid A; larger wavy line, core oligosaccharide) is
synthesized in the cytoplasm and flipped over the IM by MsbA. LptA
and LptB are part of a protein machine that transports LPS across the
periplasm to the OM. A transmembrane component (YrbK?) is pos-
tulated to complete the IM-bound ABC transporter. Additional pro-
teins (?) could be part of the complex. The Imp/RlpB complex is
thought to mediate the insertion of the newcomer LPS into the OM.
VOL. 189, 2007 CHARACTERIZATION OF E. COLI lptA AND lptB 251
upstream in the yrbG-lptB locus (Fig. 1) and predicted to be
membrane associated (35).
Two main scenarios for the transit of LPS between the IM
and the OM have been proposed: transit at sites of contact
between the IM and the OM (12, 40) and chaperone-mediated
transport through the periplasm (32). Transport of Lpp to the
OM proceeds via the latter mechanism, mediated by the
periplasmic chaperone LolA and the ABC transporter LolCDE
(41). The former mechanism has been postulated for LPS
transport based on the observation that Lpp, but not LPS,
could be released from spheroplasts in the presence of
periplasmic extracts (40). The data presented in this work do
not favor either hypothesis.
Overall, our data are consistent with the LPS biogenesis
model shown in Fig. 7. LPS synthesized in the inner leaflet of
the IM is flipped to the outer leaflet by MsbA. LPS may be
escorted through the periplasm by a dedicated chaperone ma-
chine that includes LptA, with the system energized by the IM
protein complex containing LptB and, possibly, YrbK. Imp,
RlpB, and possibly additional proteins could then function in
receiving and assembling LPS at the OM.
We are grateful to Alessandro Prinetti for his help with the Beta-
Imager analysis. We also thank Anna Villa for help with light micros-
This work was initially supported by joint grants from the “Ministero
dell’Istruzione, dell’Universita ` e della Ricerca” and the “Universita `
degli Studi di Milano” (Programmi di Rilevante Interesse Nazionale
2002 and FIRB 2001).
1. Bachmann, B. J. 1987. Derivatives and genotypes of some mutant derivatives
of Escherichia coli K12, p. 1191–1219. In J. L. Ingraham, K. B. Low, B.
Magasanik, M. Schaechter, and H. E. Umbarger (ed.), Escherichia coli and
Salmonella typhimurium: cellular and molecular biology. ASM Press, Wash-
2. Bayer, M. E. 1968. Areas of adhesion between wall and membrane of Esch-
erichia coli. J. Gen. Microbiol. 53:395–404.
3. Bayer, M. E. 1991. Zones of membrane adhesion in the cryofixed envelope
of Escherichia coli. J. Struct. Biol. 107:268–280.
4. Bishop, R. E., H. S. Gibbons, T. Guina, M. S. Trent, S. I. Miller, and C. R.
Raetz. 2000. Transfer of palmitate from phospholipids to lipid A in outer
membranes of gram-negative bacteria. EMBO J. 19:5071–5080.
5. Bos, M. P., B. Tefsen, J. Geurtsen, and J. Tommassen. 2004. Identification
of an outer membrane protein required for the transport of lipopolysaccha-
ride to the bacterial cell surface. Proc. Natl. Acad. Sci. USA 101:9417–9422.
6. Bos, M. P., and J. Tommassen. 2004. Biogenesis of the Gram-negative
bacterial outer membrane. Curr. Opin. Microbiol. 7:610–616.
7. Braun, M., and T. J. Silhavy. 2002. Imp/OstA is required for cell envelope
biogenesis in Escherichia coli. Mol. Microbiol. 45:1289–1302.
8. Chaveroche, M. K., J. M. Ghigo, and C. d’Enfert. 2000. A rapid method for
efficient gene replacement in the filamentous fungus Aspergillus nidulans.
Nucleic Acids Res. 28:e97.
9. Cherepanov, P. P., and W. Wackernagel. 1995. Gene disruption in Esche-
richia coli: TcR and KmR cassettes with the option of Flp-catalyzed excision
of the antibiotic-resistance determinant. Gene 158:9–14.
10. Dartigalongue, C., D. Missiakas, and S. Raina. 2001. Characterization of the
Escherichia coli sigma E regulon. J. Biol. Chem. 276:20866–20875.
11. Datsenko, K. A., and B. L. Wanner. 2000. One-step inactivation of chromo-
somal genes in Escherichia coli K-12 using PCR products. Proc. Natl. Acad.
Sci. USA 97:6640–6645.
12. Doerrler, W. T. 2006. Lipid trafficking to the outer membrane of Gram-
negative bacteria. Mol. Microbiol. 60:542–552.
13. Doerrler, W. T., H. S. Gibbons, and C. R. Raetz. 2004. MsbA-dependent
translocation of lipids across the inner membrane of Escherichia coli. J. Biol.
14. Doerrler, W. T., M. C. Reedy, and C. R. Raetz. 2001. An Escherichia coli
mutant defective in lipid export. J. Biol. Chem. 276:11461–11464.
15. Hanahan, D. 1983. Studies on transformation of Escherichia coli with plas-
mids. J. Mol. Biol. 166:557–580.
16. Ishidate, K., E. S. Creeger, J. Zrike, S. Deb, B. Glauner, T. J. MacAlister,
and L. I. Rothfield. 1986. Isolation of differentiated membrane domains from
Escherichia coli and Salmonella typhimurium, including a fraction containing
attachment sites between the inner and outer membranes and the murein
skeleton of the cell envelope. J. Biol. Chem. 261:428–443.
17. Kellenberger, E. 1990. The ‘Bayer bridges’ confronted with results from
improved electron microscopy methods. Mol. Microbiol. 4:697–705.
18. Kunz, D. A., and P. J. Chapman. 1981. Catabolism of pseudocumene and
3-ethyltoluene by Pseudomonas putida (arvilla) mt-2: evidence for new func-
tions of the TOL (pWWO) plasmid. J. Bacteriol. 146:179–191.
19. Maunnsbach, A. B., and B. A. Afzelius. 1999. Biomedical electron micros-
copy. Academic Press, London, United Kingdom.
20. Mecsas, J., P. E. Rouviere, J. W. Erickson, T. J. Donohue, and C. A. Gross.
1993. The activity of sigma E, an Escherichia coli heat-inducible sigma-factor,
is modulated by expression of outer membrane proteins. Genes Dev. 7:2618–
21. Meredith, T. C., P. Aggarwal, U. Mamat, B. Linder, and R. W. Woodard.
2006. Redefining the requisite lipopolysaccharide structure in Escherichia
coli. ACS Chem. Biol. 1:33–42.
22. Miller, J. H. 1992. A short course in bacterial genetics: a laboratory manual
and handbook for Escherichia coli and related bacteria. Cold Spring Harbor
Laboratory Press, Cold Spring Harbor, NY.
23. Missiakas, D., J. M. Betton, and S. Raina. 1996. New components of protein
folding in extracytoplasmic compartments of Escherichia coli SurA, FkpA
and Skp/OmpH. Mol. Microbiol. 21:871–884.
24. Missiakas, D., M. P. Mayer, M. Lemaire, C. Georgopoulos, and S. Raina.
1997. Modulation of the Escherichia coli ?E (RpoE) heat-shock transcrip-
tion-factor activity by the RseA, RseB and RseC proteins. Mol. Microbiol.
25. Oliver, D. B., and J. Beckwith. 1982. Regulation of a membrane component
required for protein secretion in Escherichia coli. Cell 30:311–319.
26. Osborn, M. J., J. E. Gander, E. Parisi, and J. Carson. 1972. Mechanism of
assembly of the outer membrane of Salmonella typhimurium. Isolation and
characterization of cytoplasmic and outer membrane. J. Biol. Chem. 247:
27. Polissi, A., and C. Georgopoulos. 1996. Mutational analysis and properties of
the msbA gene of Escherichia coli, coding for an essential ABC family
transporter. Mol. Microbiol. 20:1221–1233.
28. Raetz, C. R., and C. Whitfield. 2002. Lipopolysaccharide endotoxins. Annu.
Rev. Biochem. 71:635–700.
29. Raivio, T. L., and T. J. Silhavy. 1999. The ?E and Cpx regulatory pathways:
overlapping but distinct envelope stress responses. Curr. Opin. Microbiol.
30. Reuhs, B. L., R. W. Carlson, and J. S. Kim. 1993. Rhizobium fredii and
Rhizobium meliloti produce 3-deoxy-D-manno-2-octulosonic acid-containing
polysaccharides that are structurally analogous to group II K antigens (cap-
sular polysaccharides) found in Escherichia coli. J. Bacteriol. 175:3570–3580.
31. Reynolds, E. S. 1963. The use of lead citrate at high pH as an electron
opaque stain in electron microscopy. J. Cell Biol. 17:208–212.
32. Ruiz, N., D. Kahne, and T. J. Silhavy. 2006. Advances in understanding
bacterial outer-membrane biogenesis. Nat. Rev. Microbiol. 4:57–66.
33. Ruiz, N., and T. J. Silhavy. 2005. Sensing external stress: watchdogs of the
Escherichia coli cell envelope. Curr. Opin. Microbiol. 8:122–126.
34. Simons, R. W., F. Houman, and N. Kleckner. 1987. Improved single and
multicopy lac-based cloning vectors for protein and operon fusions. Gene
35. Sperandeo, P., C. Pozzi, G. Deho, and A. Polissi. 2006. Non-essential KDO
biosynthesis and new essential cell envelope biogenesis genes in the Esche-
richia coli yrbG-yhbG locus. Res. Microbiol 157:547–558.
36. Sprott, G. D., S. F. Kova, and C. A. Schnaitman. 1994. Cell fractionation, p.
72–103. In P. Gerhardt, R. G. E. Murray, W. A. Wood, and N. R. Krieg (ed.),
Methods for general and molecular bacteriology. ASM Press, Washington,
37. Stenberg, F., P. Chovanec, S. L. Maslen, C. V. Robinson, L. L. Ilag, G. von
Heijne, and D. O. Daley. 2005. Protein complexes of the Escherichia coli cell
envelope. J. Biol. Chem. 280:34409–34419.
38. Tam, C., and D. Missiakas. 2005. Changes in lipopolysaccharide structure
induce the ?E-dependent response of Escherichia coli. Mol. Microbiol. 55:
39. Tefsen, B., M. P. Bos, F. Beckers, J. Tommassen, and H. de Cock. 2005.
MsbA is not required for phospholipid transport in Neisseria meningitidis.
J. Biol. Chem. 280:35961–35966.
40. Tefsen, B., J. Geurtsen, F. Beckers, J. Tommassen, and H. de Cock. 2005.
Lipopolysaccharide transport to the bacterial outer membrane in sphero-
plasts. J. Biol. Chem. 280:4504–4509.
41. Tokuda, H., and S. Matsuyama. 2004. Sorting of lipoproteins to the outer
membrane in Escherichia coli. Biochim. Biophys. Acta 1694:IN1–IN9.
252SPERANDEO ET AL.J. BACTERIOL.
42. Trent, M. S., A. A. Ribeiro, S. Lin, R. J. Cotter, and C. R. Raetz. 2001. An Download full-text
inner membrane enzyme in Salmonella and Escherichia coli that transfers
4-amino-4-deoxy-L-arabinose to lipid A: induction on polymyxin-resistant
mutants and role of a novel lipid-linked donor. J. Biol. Chem. 276:43122–
43. Wu, T., A. C. McCandlish, L. S. Groenberger, S. S. Chng, T. J. Silhavy, and
D. Kahne. 2006. Identification of a protein complex that assembles lipopoly-
saccharide in the outer membrane of Escherichia coli. Proc. Natl. Acad. Sci.
44. Zhou, Z., K. A. White, A. Polissi, C. Georgopoulos, and C. R. Raetz. 1998.
Function of Escherichia coli MsbA, an essential ABC family transporter, in
lipid A and phospholipid biosynthesis. J. Biol. Chem. 273:12466–12475.
VOL. 189, 2007 CHARACTERIZATION OF E. COLI lptA AND lptB253