This information is current as
of June 14, 2017.
Lysosomal Accumulation and Recycling of
Lipopolysaccharide to the Cell Surface of
Murine Macrophages, an In Vitro and In
Vivo Study
Claire Forestier, Edgardo Moreno, Javier Pizarro-Cerda and
Jean-Pierre Gorvel
J Immunol 1999; 162:6784-6791; ;
http://www.jimmunol.org/content/162/11/6784
Subscription
Permissions
Email Alerts
This article cites 77 articles, 46 of which you can access for free at:
http://www.jimmunol.org/content/162/11/6784.full#ref-list-1
Information about subscribing to The Journal of Immunology is online at:
http://jimmunol.org/subscription
Submit copyright permission requests at:
http://www.aai.org/About/Publications/JI/copyright.html
Receive free email-alerts when new articles cite this article. Sign up at:
http://jimmunol.org/alerts
The Journal of Immunology is published twice each month by
The American Association of Immunologists, Inc.,
1451 Rockville Pike, Suite 650, Rockville, MD 20852
Copyright © 1999 by The American Association of
Immunologists All rights reserved.
Print ISSN: 0022-1767 Online ISSN: 1550-6606.
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
References
Lysosomal Accumulation and Recycling of Lipopolysaccharide
to the Cell Surface of Murine Macrophages, an In Vitro and
In Vivo Study1
Claire Forestier,* Edgardo Moreno,† Javier Pizarro-Cerda,* and Jean-Pierre Gorvel2*
L
ipopolysaccharide, the most abundant component of the
Gram-negative bacteria outer membrane, is the main Abinducing Ag during bacterial infections and one of the
most biologically active molecules (1). LPS released in the body
fluids by live or killed bacteria in the form of blebs or membrane
vesicles (2) is readily ingested by professional phagocytes following pinocytosis, receptor-mediated endocytosis, macropinocytosis,
or phagocytosis (3–7). Alternatively, phagocytosed Gram-negative
bacteria may release LPS within the intracellular milieu (8, 9).
Enterobacterial LPS has been detected in phagosomes, endosomes,
the Golgi complex, the nucleus, mitochondria, and cytoplasm (6,
10 –12). It has been shown that during the first 2 days after ingestion, LPS is mainly found in the form of small, loosely packed,
vacuoles that later seem to coalesce into larger and tightly packed
compartments (13). However, the characterization of the LPS-containing compartments during its intracellular routing in a timedependent manner has not as yet been determined.
Due to their accumulation within macrophages, LPSs have been
regarded as slowly processable molecules (14 –16). Macrophages
are able to deacylate and dephosphorylate the enterobacterial lipid
A moiety (17–19), which is eventually exocytosed to the extracel-
*Centre d’Immunologie de Marseille-Luminy, Parc Scientifique de Luminy, Case,
Marseille, France; and †Programa de Investigacion en Enfermedades Tropicales, Universidad Nacional, Heredia, Costa Rica
Received for publication November 30, 1998. Accepted for publication March
17, 1999.
The costs of publication of this article were defrayed in part by the payment of page
charges. This article must therefore be hereby marked advertisement in accordance
with 18 U.S.C. Section 1734 solely to indicate this fact.
1
This work was supported by grants from Institut National de la Santé et de la
Recherche Médicale (INSERM; 4N004B) and institutional grants from INSERM and
Centre National de la Recherche Scientifique. J.P.-C. is a recipient of a fellowship
from the Centre de la Recherche Scientifique (France), and C.F. from the Minister of
Education and Research (France).
2
Address correspondence and reprint requests to Dr. Jean-Pierre Gorvel, Centre
d’Immunologie de Marseille-Luminy, case 906, 13288 Marseille cedex 9, France.
E-mail address: [email protected]
Copyright © 1999 by The American Association of Immunologists
lular medium (20). Other investigations have suggested that the
O-chain and the core oligosaccharide of enterobacterial LPS are
also degraded within macrophages (13, 20) by mechanisms that
have not yet been elucidated. The slow processing of LPS within
phagocytic cells may be due to the absence of a molecular machinery capable of efficiently digesting it. Alternatively, LPS may
directly alter the constitutive intracellular trafficking of molecules
within cells perhaps in conjunction with its toxic effect. In this
respect, the elucidation of the trafficking of LPS and the identification of LPS-containing compartments are necessary to understand better the pathophysiologic effects mediated by this molecule
and to resolve the fate and the intracellular processing of nonprotein Ags. This last aspect is relevant in the light of investigations,
which have clearly demonstrated that microbial glycolipids may be
presented to T cells in the context of nonclassical histocompatibility Ags (21, 22).
In this work, we detailed in a time-dependent manner the intracellular trafficking of the low endotoxic Brucella abortus LPS (23)
in murine peritoneal macrophages. We studied the fate of LPS
after in vitro incubation and in vivo i.p. injection of purified LPS,
and after Brucella infection in vitro. The results show that LPS
follows the endocytic pathway described for proteins Ags, but with
unusually slow kinetics. We observed that Brucella LPS initially
accumulates within lysosomal compartments, followed by a gradual recycling to the plasma membrane, where it arranges as large
clusters. In contrast to enterobacterial LPS, no alteration in the
immunochemical properties of Brucella LPS was observed in spite
of the slow trafficking within lysosomes, suggesting that this molecule resists intracellular degradation mechanisms.
Materials and Methods
Mice and cells
Eight- to fifteen-week-old C3H/HeN female mice (Jackson ImmunoResearch, West Grove, PA) were used in all experiments. Peritoneal murine
macrophages were obtained after cervical dislocation from normal mice or
from mice injected i.p. with 0.4 ml of B. abortus S19 LPS (1 mg/ml). Each
0022-1767/99/$02.00
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
In this study, we detailed in a time-dependent manner the trafficking, the recycling, and the structural fate of Brucella abortus LPS
in murine peritoneal macrophages by immunofluorescence, ELISA, and biochemical analyses. The intracellular pathway of B.
abortus LPS, a nonclassical endotoxin, was investigated both in vivo after LPS injection in the peritoneal cavity of mice and in vitro
after LPS incubation with macrophages. We also followed LPS trafficking after infection of macrophages with B. abortus strain
19. After binding to the cell surface and internalization, Brucella LPS is routed from early endosomes to lysosomes with unusual
slow kinetics. It accumulates there for at least 24 h. Later, LPS leaves lysosomes and reaches the macrophage cell surface. This
recycling pathway is also observed for LPS released by Brucella S19 following in vitro infection. Indeed, by 72 h postinfection,
bacteria are degraded by macrophages and LPS is located inside lysosomes dispersed at the cell periphery. From 72 h onward,
LPS is gradually detected at the plasma membrane. In each case, the LPS present at the cell surface is found in large clusters with
the O-chain facing the extracellular medium. Both the antigenicity and heterogenicity of the O-chain moiety are preserved during
the intracellular trafficking. We demonstrate that LPS is not cleared by macrophages either in vitro or in vivo after 3 mo, exposing
its immunogenic moiety toward the extracellular medium. The Journal of Immunology, 1999, 162: 6784 – 6791.
The Journal of Immunology
killed mouse received an i.p. injection of 20 ml of sterile PBS/10% sucrose
at 15°C. After injection, the abdomen of animal was massaged and the
liquid was extracted from the peritoneal cavity. Pooled fluids were centrifuged at 1200 rpm for 10 min at 4°C and the pellets were resuspended in
DMEM (Life Technologies, Cergy-Pontoise, France) supplemented with
10% FCS, 2 mM glutamine, 10 mM sodium pyruvate, 10 mM nonessential
amino acids, 10 mM HEPES, 100 U/ml penicillin, and 100 mg/ml streptomycin (all from Life Technologies). A total of 2 3 105 peritoneal cells
was plated on 12-mm glass coverslips in 24-well tissue culture plastic
dishes, at 106 cells/ml in 96-well plates (Costar, Cambridge, MA) or at 106
cells/35-mm dish (Nunc, Poly Labo) for immunofluorescence, ELISA, or
immunoblotting analysis, respectively. After incubation for 1 h at 37°C
under 5% CO2, nonadherent cells were removed from the wells by aspiration, and the adherent macrophages were rinsed twice with PBS and
incubated with fresh culture medium.
Bacteria and LPS preparations
Cell infection and LPS endocytosis
After removal of culture medium, macrophages grown on coverslips in
24-well tissue plates were inoculated with 500 ml of a bacterial suspension
in culture media at a rate of 50 bacteria/cell. Culture plates were centrifuged for 10 min at 1000 rpm at room temperature, followed by incubation
for 20 min at 37°C under 5% CO2. Cells were washed five times with
culture medium to remove nonadherent bacteria, and monolayers were further incubated with culture medium containing 50 mg/ml of gentamicin
(Sigma) to kill extracellular brucellae. In long-term experiments, this medium was replaced twice: at 1 h with medium containing 25 mg/ml gentamicin, and at 24 h with medium supplemented with 5 mg/ml gentamicin.
Macrophage layers grown on glass coverslips (for immunofluorescence
experiments), 96-well plates (for ELISA), or 35-mm plastic dishes (for
immunoblotting analysis) were pulsed with B. abortus LPS. In each experiment, 15 mg/ml of LPS was added for 1 h at 4°C, followed by extensive
washings with culture media. Then macrophages were incubated in cell
culture medium at 37°C for different time intervals (10 min to 48 h) and
processed for immunofluorescence, ELISA, or immunoblotting analysis.
Antibodies
Murine mAb against O-chain C/Y epitope (Baps C/Y) was produced and
characterized, as previously described (29). Cow polyclonal serum against
B. abortus S-LPS3 was described in earlier works (29). Briefly, the serum
was collected from a B. abortus-infected cow and was shown to react
strongly against two O-chain epitopes (C/Y and A) from S-LPS, core
epitopes from isolated R-LPS, and isolated lipid A epitopes. At the dilution
used, this serum does not react against core or lipid A when LPS molecule
is complete (core and lipid A epitopes are hidden due to the presence of
3
Abbreviations used in this paper: S-LPS, smooth LPS; R-LPS, rough LPS; CIM6PR, cation-independent mannose-6-phosphate receptor.
4
C. Forestier, E. Moreno, A. Phalipon, P. Sansonetti, and J.-P. Gorvel. Interactions of
Brucella LPS and MHC class II molecules. Submitted for publication.
5
W. Beatty, S. Méresse, J. Davoust, P. Gounon, P. Sansonnetti, and J. P. Gorvel.
Trafficking of Shigella LPS in polarized intestinal epithelial cells. J. Cell Biol. In
press.
O-chain). Rabbit polyclonal antisera against R-LPS were produced by immunizing rabbits with a purified preparation of B. abortus 45/20 R-LPS
(30). This antiserum reacts against core epitopes from the R-LPS, but does
not react against the O-chain, core epitope from S-LPS, and lipid A (29).
Rabbit IgGs anti-cow Ig and Baps C/Y mAb were linked to horseradish
peroxidase, as previously described (31). At the dilution used, none of the
polyclonal or mAbs used demonstrated reactivity against LPS-untreated
macrophages. Rabbit polyclonal IgGs anti-cation-independent mannose-6phosphate receptor (anti-CI-M6PR; Dr. B. Hoflack, Institut Pasteur de
Lille, Lille, France); rabbit anti-cathepsin D (Dr. S. Kornfeld, Washington
University School of Medicine, St. Louis, MO); and rabbit anti-giantin (Dr.
H. P. Hauri, University of Basel, Basel, Switzerland) were used for immunofluorescence experiments. Secondary Abs used were: goat Texas redconjugated anti-cow IgG and donkey FITC-conjugate anti-rabbit IgG
(Jackson ImmunoResearch). Rabbit IgG-peroxidase anti-mouse Ig and goat
IgG-peroxidase anti-rabbit Ig conjugates were used (Sigma) as secondary
Abs for Western blotting and ELISA.
Confocal microscopy
For fluorescent confocal microscopy, macrophages grown in coverslips
were fixed at room temperature with 3.7% paraformaldehyde in PBS, pH
7.4, for 20 min, followed by 10-min incubation with 0.1 M glycine to
saturate aldehyde groups and with 0.1% saponin for cell permeabilization.
Then cells were incubated with a PBS solution containing 0.1% saponin/5% mouse serum/5% horse serum for 20 min to block unspecific binding. Primary Abs diluted in the same buffer were added to the cells for 30
min. After extensive washings with 0.05% saponin in PBS, macrophages
were incubated for 30 min with secondary fluorescent Abs. Coverslips with
adherent macrophages were washed, mounted in Mowiol (Hoechst,
Frankurt, Germany), and viewed under a Leica TCS 4D confocal microscope (Leica Lasertechnik, Heildeberg, Germany). In all immunofluorescence experiments, LPS was revealed using the antiserum from infected
cow, followed by anti-cow IgG Ab conjugated to fluorescein.
Western blotting
After internalization of B. abortus LPS, macrophages were lysed in
PBS/1% Nonidet P-40 detergent for 30 min, and treated with 1 mg/ml of
proteinase K (Sigma) at 37°C for 30 min. Lysates were run in 12% SDSPAGE and electrophoretically transferred onto Immobilon-P membranes
(Millipore, Bedford, MA). Membranes were blocked in PBS/5% milk/
0.05% Tween-20 (Sigma) for 2 h. Primary and secondary Abs were successively added in this buffer, each being left for 2 h before staining with
the enhanced chemoluminescence system (ECL; Amersham, Courtaboeuf,
France). As primary anti-LPS Abs, Baps C/Y, cow, and rabbit antisera
were used to detect the different epitopes of LPS in macrophage cell lysates.
Dot-blot analysis
Peritoneal liquids from LPS-inoculated mice were recovered by injecting 5
ml of PBS/10% sucrose in peritoneal cavity; the liquid was extracted and
centrifuged to eliminate macrophages. Supernatants were filtered through a
0.45-mm cutoff membrane (Gelman Sciences, Ann Arbor, MI) to eliminate
cellular debris. A total of 100 ml of supernatant was applied to Immobilon-P membrane (Millipore) in a dot-blot manifold (Bio-Rad, Hercules,
CA). The membrane was removed from the manifold and blocked with
0.2% BSA for 1 h before adding the peroxidase-conjugated anti-O-chain
Ab (Baps C/Y). Detection has been done with the enhanced chemoluminescence system.
ELISA assays
After LPS internalization, macrophages were fixed with 3.7% paraformaldehyde in PBS, then incubated with PBS/10% mouse serum/0.1% saponin
to block unspecific sites and to permeabilize the cells. Parallel cultures
were not permeabilized to measure LPS exclusively on the cell surface.
Cells were then incubated with 100 ml of the antiserum from infected cow
diluted in PBS/5% normal mouse serum, with or without the addition of
saponin for nonpermeabilized and permeabilized samples, respectively.
After 45-min incubation, cells were washed extensively with PBS, and 100
ml of horseradish peroxidase-conjugated anti-bovine Ig was added for 45
min at room temperature. After washings, 100 ml of tetramethylbenzidine
substrate (Sigma) was added to the cells. After 30-min incubation, the
enzymatic reaction was stopped by adding 10 ml of 0.5 M H2SO4, and the
color reaction was read at 450 nm OD. To estimate at which LPS dose the
OD corresponded, various fixed amounts of LPS were deposited on an
Immobilon-P membrane in a dot-blot manifold, as described above. Each
spot was cut off, and the membrane was then deposited in distinct wells on
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
The characteristics of smooth type B. abortus S19 (Professional Biological,
Denver, CO) have been previously described (24). Bacteria were grown at
37°C in tryptic soy broth (TSB; Difco, Detroit, MI) to stationary phase, and
aliquots were frozen at 270°C in TSB-glycerol 30%. A log-phase culture
bacteria was prepared by incubating a thawed aliquot (5 3 1010 CFU/ml)
in TSB for 15– 48 h at 37°C under agitation. Bacterial numbers were determined by comparing the OD at 600 nm with a standard curve.
The extraction and purification of LPS from smooth and rough B. abortus have been described elsewhere (25, 26). Briefly, smooth Brucella LPS
was obtained from the phenolic phase of phenol-water extraction method
(27). Purification of LPS (20 mg/ml LPS in 5 mM MgCl2/10 mM Tris-HCl,
pH 7.5) was conducted by repeated digestion with nucleases and proteinase
K (Sigma, St. Louis, MO) for 24 h at 25°C. The LPS was then centrifuged
at 100,000 3 g for 6 h at 4°C, and the removal of phospholipids and
ornithine lipids was achieved with chloroform/methanol/water (1:8:0.8,
v/v/v), followed by chloroform/methanol/7 M ammonia (65:25:4, v/v/v).
LPS was then reextracted with phenol/water and recovered by centrifugation at 100,000 3 g for 6 h at 4°C. All LPS preparations were lyophilized
after extensive dialysis and analyzed by standard procedures (26, 28). For
internalization studies and injections in mice, LPS was dissolved in water
at the appropriate concentration with the aid of sonication and autoclaved
before use.
6785
6786
24-well plates. The amount of LPS was revealed by ELISA assays (described above) using peroxidase-conjugated anti-O-chain Ab (Baps C/Y).
Results
LPS is delivered with slow kinetics to lysosomes, where it
accumulates
To define the internalization pathway of Brucella LPS in macrophages, we compared, by indirect immunofluorescence, the intracellular distribution of LPS with several markers of endocytic
compartments and the Golgi apparatus. Macrophages were initially
incubated with LPS for 1 h at 4°C, followed by different periods of
chase at 37°C. After 10 min, a punctate LPS staining pattern, located intracellularly at the cell periphery, colocalized with FITCtransferrin, indicating that LPS had reached the early endosomal
network (Fig. 1, A and B). From early endosomes, internalized
material can be recycled back to the plasma membrane or directed
to more acidic intracellular compartments, such as late endosomes
and eventually lysosomes (32, 33). From 15 min onward, LPS
localized mostly within FITC-transferrin-negative vacuoles, suggesting a transient and rapid passage of LPS through early endocytic organelles (Fig. 1, C and D). After 90 min, LPS became
concentrated in intracellular compartments positive for the CIM6PR (Fig. 1, E and F). This marker has been described as cycling
between the Golgi apparatus and late endosomes and, depending
on cell type, it accumulates either in the trans-Golgi network or in
FIGURE 2. Transient accumulation of Brucella LPS in lysosomes. In A,
macrophages untreated with LPS were stained with the antiserum from a B.
abortus-infected cow (Aa) and with anti-cathepsin D Ab (Ab). In B, macrophages were incubated with LPS as in Fig. 1. Times of chase were from
1– 48 h. LPS, detected with the antiserum from Brucella-infected cow, is
shown in Ba, Bc, and Be, and cathepsin D in Bb, Bd, and Bf. After 5 h at
37°C, LPS (Ba) reached lysosomes positive for cathepsin D (Bb) and accumulated in this compartment even after 24 h (Bc and Bd). At 48 h, LPS
(Be) was mainly found outside lysosomes (Bf). Bar 5 10 mm.
late endosomes (34). Since no colocalization was observed with
giantin, a specific marker of the Golgi apparatus (Fig. 1, G and H),
we concluded that LPS was transferred from early to late endosomes. From late endosomes, LPS took about 5 h to reach cathepsin D-positive lysosomal compartments (Fig. 2, Ba and Bb). From
this time up to 24 h, LPS steadily accumulated within the lysosomes (Fig. 2, Bc and Bd), demonstrating that LPS is not eliminated from the host cell. The slow kinetics of LPS trafficking
seems to be independent of the activation state of macrophages,
since in macrophages that were pretreated with IFN-g for 48 h
before LPS addition, LPS reached the lysosomes after 5-h internalization (data not shown).
LPS escapes from lysosomes and recycles to the plasma
membrane
The persistence of LPS in macrophages prompted us to investigate
the fate of LPS at late time points. At 48 h, LPS was still present
in macrophages. Although a minor proportion of LPS-containing
vesicles remained positive for the lysosomal marker cathepsin D,
the majority of LPS was found within intracellular nonlysosomal
compartments (Fig. 2, Be and Bf) and at the plasma membrane
forming large clusters (Fig. 3B). These results suggest that, after
lysosomal accumulation, LPS escaped lysosomes and migrated to
the cell surface. We also estimated the relative amounts of cell
surface-bound versus intracellular LPS by ELISA (Fig. 3A). Even
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
FIGURE 1. Intracellular trafficking of Brucella LPS in murine peritoneal macrophages. Cells were incubated with 15 mg/ml of LPS in cell
culture medium for 1 h at 4°C, then washed and further incubated for
various times at 37°C. After paraformaldehyde fixation, macrophages were
processed for double immunofluorescence. LPS was detected by indirect
immunofluorescence using the antiserum from Brucella-infected cow. After 10 min of internalization at 37°C, LPS (A) colocalized with transferrinFITC (B) and, at 60 min, LPS (C) was no longer associated with transferrin-FITC (D). At 90 min, LPS (E, G) was found in a compartment
positive for the CI-M6PR (F) and negative for giantin, a specific marker of
the Golgi apparatus (H). Bar 5 10 mm.
Brucella abortus LPS TRAFFICKING IN MACROPHAGES
The Journal of Immunology
6787
FIGURE 3. Recycling of Brucella LPS to the cell surface of macrophages. In A and B, peritoneal macrophages were harvested from normal
mice, plated for 2 h (5 3 105 cells/ml), then incubated with Brucella LPS
(15 mg/ml) for 1 h at 4°C. In C and D, macrophages were harvested from
LPS-injected mice, at 60 days postinjection. After several washings, cells
were incubated at 37°C and fixed. One-half of the samples were permeabilized with saponin. Quantification of intracellular versus cell surfacebound LPS was analyzed by ELISA assay (A, C) using the antiserum from
a B. abortus-infected cow. Nonpermeabilized cells allowed the quantification of cell surface-associated LPS (f), whereas permeabilization gave the
measurements of total LPS (intracellular 1 surface-bound) (E). The
amount of intracellular LPS (L) was calculated by the subtraction of the
values obtained with nonpermeabilized samples from those obtained with
permeabilized samples. Immunofluorescence (B, D) shows the distribution
of Brucella LPS in permeabilized (upper panel) and nonpermeabilized
macrophages (lower panel), following 48 h (B) or 2 h (C) of chase in vitro.
f, Surface-bound LPS; L, total LPS; E, intracellular LPS.
peritoneal fluids occurred between 17 and 20 days (not shown),
and inoculated LPS internalized by resident macrophages remained associated with these cells for at least 3 mo after injection
(Fig. 4). Similarly to what was observed in cell cultures after 2
days, LPS aggregates forming large clusters, distributed inside and
outside lysosomal compartments as well as at the plasma membrane (Figs. 4 and 3D). This unique cellular distribution pattern
persisted up to 90 days postinjection (not shown). Peritoneal macrophages from LPS-injected mice, harvested at 60 days postinjection, were then cultured in vitro in the absence of LPS. Fig. 3C
shows that LPS started to recycle from lysosomal and nonlysosomal compartments to the plasma membrane after 48 h. The majority of LPS molecules reached the macrophage cell surface after
5 days (Fig. 3C), confirming the results shown in Fig. 3A.
The immunochemical properties of Brucella LPS are unaltered
during its intracellular trafficking
though the majority of LPS was rapidly internalized after 10 min,
as shown in Fig. 1, A and B, after 2 h, 25% of total LPS still resided
at the plasma membrane (Fig. 3A). Complete internalization occurred by 24 h. Then LPS gradually reached the macrophage cell
surface, and at 48 h, most of the LPS was detected by anti-O-chain
Abs, at the external leaflet of the plasma membrane (Fig. 3A). We
evaluated the total amount of macrophage-associated LPS to be 1
ng of LPS/106 cells (see ELISA assays in Materials and Methods).
In addition, the total amount of LPS associated with macrophages
did not vary significantly during the in vitro chase periods, and we
never detected exocytosed LPS in the cell culture supernatant by
dot-blot analysis (not shown). Altogether, these data support the
idea that LPS is not eliminated by macrophages.
As shown in Fig. 5, A and B, native Brucella S-LPS displays a
characteristic m.w. heterogeneity related to the different lengths of
the O- chain, core oligosaccharide, and lipid A (35), whereas RLPS appears as a lower m.w. band (Fig. 5C). After internalization
and trafficking within macrophages, LPS did not show any significant SDS-PAGE profile variation or altered antigenic reactivity
against a collection of monoclonal anti-O-chain Abs (Fig. 5A) as
well as polyclonal Abs from a B. abortus-infected cow, and was
shown to react strongly against two O-chain epitopes (C/Y and A)
from S-LPS, core epitopes from isolated R-LPS, and isolated lipid
A epitopes (Fig. 5B), suggesting that LPS was not cleaved during
its intracellular trafficking in peritoneal macrophages. In addition,
we never detected the appearance of R-LPS (Fig. 5C), indicating
that the O-chain moiety was not cleaved from S-LPS. By comparing the signal intensity corresponding to cell-associated LPS with
that of purified LPS, the amount of cell-associated LPS was estimated to 1 ng/106 cells in agreement with ELISA measurements.
Brucella LPS remains associated with peritoneal macrophages
for several months
Intracellular LPS released from digested bacteria is detected at
the plasma membrane
To investigate the fate of LPS in vivo, we injected C3H/HeN mice
i.p. with 400 mg of Brucella LPS. The kinetics of LPS removal
from the peritoneal cavity was followed at various times after LPS
injection by dot-blot analysis. Complete clearance of LPS from the
We have shown that the attenuated B. abortus S19 strain is degraded within lysosomes of phagocytes (36). To explore the fate of
LPS released intracellularly after the degradation of phagocytosed
attenuated S19 bacteria in lysosomes, we followed the intracellular
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
FIGURE 4. Distribution of Brucella LPS in macrophages after peritoneal injection of LPS. Mice were i.p. injected with Brucella LPS (400 mg).
Peritoneal macrophages were recovered after 6 or 60 days and processed
for double immunofluorescence. LPS (upper panels), revealed by the
antiserum from a Brucella-infected cow, colocalizes partially even after
60 days with cathepsin D-positive compartments (lower panels).
Bar 5 10 mm.
6788
trafficking of S19 Brucella LPS within macrophages at various
times postinfection. Forty-eight hours after inoculation, most intact
bacteria were found to be within compartments positive for the
lysosomal marker cathepsin D (Fig. 6A). At 72 h, most bacteria
were degraded within lysosomes, and bacterial debris containing
LPS were found within cathepsin D-positive vesicles, scattered
throughout the cytoplasm (Fig. 6A), indicating that while bacteria
underwent enzymatic degradation in the cathepsin D-positive compartment, the epitopic structure of LPS resisted the intracellular
degradation processes. From 72 h onward, LPS was gradually detected at the macrophage plasma membrane (Fig. 6B). Finally, at
6 days postinfection, almost all infected cells exposed high levels
of LPS at their plasma membrane (Fig. 6B), indicating that a considerable proportion of glycolipids released by the degraded bacteria followed the recycling pathway described above.
Discussion
Macrophages are specialized cells for capturing, processing, and
presenting exogenous Ags to T lymphocytes. They are also the
principal target for many intracellular Gram-negative bacteria and
their endotoxins. The biological action of LPS on macrophages has
been studied extensively (37). Among the less well-characterized
events following LPS interaction with macrophages are the intracellular trafficking and processing of internalized LPS. A comprehension of these steps is critical for the understanding of the biological activity and immunogenicity of this nonpeptidic Ag.
In this study, we demonstrate that, in conditions that favor receptor-mediated endocytosis in vitro (38), LPS is rapidly internalized in early endosomes and transported, with slow kinetics, to late
endosomes and then to lysosomes, where it accumulates. Later,
LPS is found in cathepsin D-negative compartments before it
reaches the cell surface, where it forms large clusters. In contrast
to other reports, we did not detect Brucella LPS either in the Golgi
apparatus, nucleus, mitochondria, nor free in the cytoplasm (6, 10).
This discrepancy may be explained by the different experimental
procedures, including the use of different target cells and high
doses of cytotoxic LPS. Despite the fact that Brucella LPS induces
many biological effects, characteristics of so-called typical LPS
molecules, it possesses very low toxicity (23), which favors longterm studies in cultured cells and in animals (39). In general, previous studies were performed by using continuous enterobacterial
LPS incubation at 37°C, which may allow receptor-mediated uptake, macropinocytosis, and fluid-phase endocytosis (6, 10). Recent studies from Kitchens et al. demonstrated that the route and
kinetics of LPS trafficking depend upon its internalization pathway
(CD14 dependent or independent) and its aggregation state, respectively (40, 41), thus offering an explanation for the divergence
of the results obtained between different studies.
It has been proposed that the kinetics of intracellular trafficking
of different substances is independent of the nature of the ingested
molecule in many respects (42– 44). However, several studies have
demonstrated that LPS follows the endocytic pathway with slow
kinetics (11) (Beatty et al., submitted). In agreement with these, we
observed a substantial delay in the intracellular transit of LPS from
early endosomes to lysosomes, in comparison with proteins such
as BSA-FITC, which is rapidly transported from early endosomes
to lysosomes within 60 min after internalization and remains located in perinuclear lysosomes until its degradation (45). The slow
kinetics of Brucella LPS traffic considerably differs from Brucella
organisms. In the case of virulent bacteria, it is known that bacteria
evade lysosomes and replicate within the endoplasmic reticulum.
For nonvirulent Brucella, the bacteria are found within phagosomes that rapidly fuse with lysosomes, where degradation occurs
12 h postinfection (36). In the latter case, Brucella LPS is consequently released from the outer membrane of the killed bacteria
and remains within lysosomes for at least 140 h. This last observation is in agreement with several investigations that have demonstrated the appearance and retention of free LPS within host
cells after phagocytosis of different bacteria (8, 9, 13, 46). It is then
possible that LPS remains in a nonfunctional lysosomal compartment in which resident proteins, such as cathepsin D and LAMP
(lysosome-associated membrane protein), have been degraded or
removed. In favor of this is the finding that indigestible polystyrene particles are contained within nonfunctional lysosomes characterized by the absence of hydrolases (44). In addition, it may be
that LPS prevents the acquisition of newly synthesized hydrolases
arriving from Golgi-derived vesicles, the consequence of which
would be the generation of hydrolase-defective lysosomes (47).
Therefore, the retention process and coalescence of LPS in large
vacuolar aggregates within macrophages seem to be a property of
this complex molecule and independent of the bacteria from which
it originates.
The clustering of LPS molecules and the formation of discrete
patches on the macrophage plasma membrane are consistent with
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
FIGURE 5. Brucella LPS analysis by immunoblotting. Cells were incubated with Brucella S-LPS (15 mg/ml) at 4°C for 1 h, washed, and
further incubated with cell culture medium at 37°C for 0.5, 2, 5, 24, 30, and
48 h. Then macrophages were lysed and the lysates were loaded on 12%
SDS-polyacrylamide gels, and after transfer to Immobilon membranes,
LPS was revealed in A using the Baps C/Y peroxidase-conjugated mAb
that recognizes specifically C/Y sugar epitopes of the O-chain moiety. In B,
LPS was revealed using a serum from a B. abortus-infected cow that recognizes the O-chain C/Y most abundant epitope and the A epitope of
S-LPS, the outer core epitope exposed in isolated R-LPS and the backbone
disaccharide of the isolated lipid A. In D, the presence of both R-LPS was
tested by using a rabbit antiserum against B. abortus 45/20 R-LPS recognizing specifically the core moiety of R-LPS. A total of 10 ng of purified
S-LPS (S) (A and B) and 100 ng of purified R-LPS (R) was deposited and
revealed, as described above.
Brucella abortus LPS TRAFFICKING IN MACROPHAGES
The Journal of Immunology
6789
FIGURE 6. Distribution of Brucella LPS in macrophages following Brucella S19 infection. Macrophages
were infected with B. abortus strain 19 for 10 min,
washed, incubated for various times at 37°C, then processed for double immunofluorescence. Infected macrophages were permeabilized with saponin (A) and labeled with both anti-cathepsin D Ab and B. abortusinfected cow serum, which detects either the LPS
associated to bacteria or LPS liberated from degraded
bacteria. In B, macrophages were not permeabilized to
monitor the presence of surface-bound LPS at different
postinfection times (48, 72, and 120 h).
the fact that LPS possesses structural and functional similarities
with sphingolipids such as ganglosides and cerebrosides (58, 59),
LPS does not recycle by the same mechanism since we never detected this molecule in the Golgi apparatus. It has also been demonstrated that processed proteic Ags recycle from late endocytic
compartments to the cell surface by transiting within specialized
MHC class II (MHC-II)-enriched compartments, where peptides
associate with Ag-presenting molecules (60). It is possible that
Brucella LPS, which is also found in late endocytic compartments,
recruits and modifies MHC-II products within lysosomes, as suggested by previous experiments in which this molecule was found
to interact and induce the formation of SDS-resistant forms (C
forms) of MHC-II proteins in B lymphocytes (61) (Forestier et al.,
submitted). The coalescence of LPS in discrete patches at the
plasma membrane may be the result of a combination of the intrinsic properties of LPS, which promotes its own arrangement as
crystal layers in the outer leaflet of membranes (62), and the transport of LPS to specific plasma membrane domains by carrier
molecules.
Macrophages are capable of processing enterobacterial LPS by
a slow mechanism that may involve a combination of oxidative
and enzymatic hydrolytic procedures (18 –20). These cells modify
the chemical structure of any of the three enterobacterial LPS moieties (lipid A, core, and O-chain) and, as a consequence, alter the
electrophoretic mobility of this molecule and its reactivity against
specific Abs (13). For instance, deacylation and dephosphorylation
of LPS result in reduced amounts of lipid A fatty acids, less binding of SDS, and slower migration in SDS-PAGE (13, 46, 63).
Oxidation and hydrolysis of core sugars diminish the imunoreactivity against core determinants and result in free lipid A detected
with specific Abs (13, 35, 64). Commonly, the resulted free Ochain is unable to migrate in SDS-PAGE due to the absence of
SDS binding sites (28, 35). Removal of sugar substituents or repeating units from the O-chain causes faster aberrant migration in
SDS-PAGE and reduced reactivity to Abs (13, 65). In spite of its
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
studies describing the preferential association of LPS with particular classes of membrane lipids (48) at specific membrane domains
(5, 49). Based on recent experiments conducted in our laboratory
in which the membrane fluidity of Brucella LPS-treated cells has
been measured (unpublished results), and experiments demonstrating the direct (50, 51) or LPS binding protein (LBP)-mediated (52,
53) intercalation of LPS into bilayers, we propose that Brucella
LPS is inserted into the cell membranes by its lipid A moiety,
exposing its immunogenic O-chain moiety toward the extracellular
medium. The existence and the fate of these LPS large clusters in
the plasma membrane remain unknown. Although it was found at
the cell surface, no free Brucella LPS was detected in supernatants
of cell cultures, as demonstrated by the ELISA and colocalization
experiments, respectively, suggesting that there is no exocytosis of
LPS from macrophages. The fact that LPS disappeared from peritoneal fluids of mice only 3 wk after its injection suggests that
phagocytosis of high doses of LPS by peritoneal cells may be the
limiting factor for its clearance in vivo. The presence of conspicuous large aggregates in peritoneal macrophages after 6 and 60
days highlights the remarkable stability of this molecule.
After storage in lysosomes, LPS escapes these compartments
and appears on the plasma membrane of macrophages, in agreement with the results previously obtained by Korn et al. (54). Although the mechanism by which LPS recycles from intracellular
compartments to the plasma membrane remains to be characterized, it is likely that LPS-containing vesicles ensure the transport
of LPS from intracellular compartments to the plasma membrane
by specific target mechanisms. It may be that the insertion of the
lipid A into the membrane of the LPS-containing compartment
recruits constitutive recycling molecules, promoting its own transportation to the cell surface. Recycling pathways have been partially characterized for complex membrane lipids or exogenous
protein Ags. For instance, ceramide-containing sphingolipids have
been shown to be translocated from the Golgi apparatus to the
plasma membrane by a vesicle-mediated process (55–57). Despite
Brucella abortus LPS TRAFFICKING IN MACROPHAGES
long residence inside lysosomes, Brucella LPS seems to remain
uncleaved. This observation is based on the unaltered migration
pattern in SDS-PAGE of the Brucella LPS extractable from macrophages, its consistent reactivity against monoclonal and polyclonal Abs, and the absence of detectable free core or lipid A
determinants. The resistance of Brucella LPS to macrophage processing may simply reflect the absence of a cellular machinery
capable of oxidizing and hydrolyzing this complex molecule. Indeed, the O-chain, the core oligosaccharide, and the lipid A dissacharide backbone of Brucella LPS are respectively more resistant to oxidation, acid hydrolysis (35, 66), and deacylation (26)
than enterobacterial LPS. In conclusion, the unique chemical structure of Brucella LPS may not be prone to regular processing mechanisms described in the case of enterobacterial LPS.
Finally, our results indicate that LPS is retained in an immunologically detectable form in murine peritoneal macrophages for
long periods of time, then recycles and associates as clusters at the
plasma membrane without detectable modification. The long permanence of LPS within cells of individuals may result at least in
several pathologic effects, such as reactive arthritis or hypersensitivity reactions, developed after acute and chronic bacterial infections (67– 69). The presence of LPS at the plasma membrane of
APC may lead to the activation of T lymphocytes, as it has been
shown for other glycolipids (70 –75). The recycling phenomenom
may explain how LPS activates some T lymphocytes, as recently
shown by others (75–78).
pathogen-derived T-cell-independent antigens on protein presentation to T cells.
Clin. Immunol. Immunopathol. 85:1.
Munford, R. S., and C. L. Hall. 1985. Uptake and deacylation of bacterial lipopolysaccharides by macrophages from normal and endotoxin-hyporesponsive
mice. Infect. Immun. 48:464.
Munford, R. S., and C. L. Hall. 1986. Detoxification of bacterial lipopolysaccharides (endotoxins) by a human neutrophil enzyme. Science 234:203.
Peterson, A. A., and S. Munford. 1987. Dephosphorylation of the lipid A moiety
of Escherichia coli lipopolysaccharide by mouse macrophages. Infect. Immun.
55:974.
Fox, E. S., P. Thomas, and S. A. Broitman. 1989. Clearance of gut-derived endotoxins by the liver: release and modification of 3H,14C-lipopolysaccharide by
isolated rat Kupffer cells. Gastroenterology 96:456.
Porcelli, S. A., C. T. Morita, and R. L. Modlin. 1996. T-cell recognition of
non-peptide antigens. Curr. Opin. Immunol. 8:510.
Melian, A., E. M. Beckman, S. A. Porcelli, and M. B. Brenner. 1996. Antigen
presentation by CD1 and MHC-encoded class I-like molecules. Curr. Opin. Immunol. 8:82.
Rasool, O., E. Freer, E. Moreno, and C. Jarstrand. 1992. Effect of Brucella abortus lipopolysaccharide on oxidative metabolism and lysozyme release by human
neutrophils. Infect. Immun. 60:1699.
Sola-Landa, A., J. Pizarro-Cerda, M.-J. Grillo, E. Moreno, I. Moriyon, J.-M.
Blasco, J.-P. Gorvel, and I. Lopez-Goni. 1998. A two-component regulatory system playing a critical role in plant pathogens and endosymbionts is present in
Brucella abortus and controls cell invasion and virulence. Mol. Microbiol. 29:
125.
Moreno, E., M. W. Pitt, L. M. Jones, G. G. Schurig, and D. T. Berman. 1979.
Purification and characterization of smooth and rough lipopolysaccharides from
Brucella abortus. J. Bacteriol. 138:361.
Moreno, E., E. Stackebrandt, M. Dorsch, J. Wolters, M. Busch, and H. Mayer.
1990. Brucella abortus 16S rRNA and lipid A reveal a phylogenetic relationship
with members of the a-2 subdivision of the class Proteobacteria. J. Bacteriol.
172:3569.
Baker, P. J., and J. B. Wilson. 1965. Chemical composition and biological properties of the endotoxin of Brucella abortus. J. Bacteriol. 90:895.
Aragon, V., R. Diaz, E. Moreno, and I. Moriyon. 1996. Characterization of Brucella abortus and Brucella melitensis native haptens as outer membrane O-type
polysaccharides independent from the smooth lipopolysaccharide. J. Bacteriol.
178:1070.
Rojas, N., E. Freer, A. Weintraub, M. Ramirez, S. Lind, and E. Moreno. 1994.
Immunochemical identification of Brucella abortus lipopolysaccharide epitopes.
Clin. Diagn. Lab. Immunol. 1:206.
Moreno, E., L. M. Jones, and D. T. Berman. 1984. Immunochemical characterization of rough Brucella lipopolysaccharide. Infect. Immun. 43:779.
Ramirez, P., J. A. Bonilla, E. Moreno, and P. Leon. 1983. Electrophoretic transfer
of viral proteins to nitrocellulose sheets and detection with peroxidase-bound
lectins and protein A. J. Immunol. Methods 62:15.
Gruenberg, J., and K. E. Howell. 1989. Membrane traffic in endocytosis: insights
from cell-free assays. Annu. Rev. Cell Biol. 5:453.
Kornfeld, S., and I. Mellman. 1989. The biogenesis of lysosomes. Annu. Rev. Cell
Biol. 5:483.
Griffiths, G., B. Hoflack, K. Simons, I. Mellman, and S. Kornfeld. 1988. The
mannose 6-phosphate receptor and the biogenesis of lysosomes. Cell 52:329.
Freer, E., N. Rojas, A. Weintraub, A. A. Lindberg, and E. Moreno. 1995. Heterogeneity of Brucella abortus lipopolysaccharides. Res. Microbiol. 146:569.
Pizarro-Cerda, J., E. Moreno, V. Sanguedolce, J. L. Mege, and J. P. Gorvel. 1998.
Virulent Brucella abortus prevents lysosome fusion and is distributed within
autophagosome-like compartments. Infect. Immun. 66:2387.
Rietschel, E. T., H. Brade, O. Holst, L. Brade, S. Muller-Loennies, U. Mamat,
U. Zahringer, F. Beckmann, U. Seydel, K. Brandenburg, et al. 1996. Bacterial
endotoxin: chemical constitution, biological recognition, host response, and immunological detoxification. Curr. Top. Microbiol. Immunol. 216:39.
Wileman, T., C. Harding, and P. Stahl. 1985. Receptor-mediated endocytosis.
Biochem. J. 232:1.
Moreno, E., D. T. Berman, and L. A. Boettcher. 1981. Biological activities of
Brucella abortus lipopolysaccharides. Infect. Immun. 31:362.
Kitchens, R. L., and R. S. Munford. 1998. CD14-dependent internalization of
bacterial lipopolysaccharide (LPS) is strongly influenced by LPS aggregation but
not by cellular responses to LPS. J. Immunol. 160:1920.
Kitchens, R. L., P.-Y. Wang, and R. S. Munford. 1998. Bacterial lipopolysaccharide can enter monocytes via two CD14-dependent pathways. J. Immunol.
161:5534.
Ward, D. M., R. Ajioka, and J. Kaplan. 1989. Cohort movement of different
ligands and receptors in the intracellular endocytic pathway of alveolar macrophages. J. Biol. Chem. 264:8164.
Ward, D. M., and J. Kaplan. 1990. The rate of internalization of different receptor-ligand complexes in alveolar macrophages is receptor-specific. Biochem. J.
270:369.
Oh, Y. K., and J. A. Swanson. 1996. Different fates of phagocytosed particles
after delivery into macrophage lysosomes. J. Cell Biol. 132:585.
Tassin, M. T., T. Lang, J. C. Antoine, R. Hellio, and A. Ryter. 1990. Modified
lysosomal compartment as carrier of slowly and non-degradable tracers in macrophages. Eur. J. Cell Biol. 52:219.
Duncan, R. L. Jr., J. Hoffman, V. L. Tesh, and D. C. Morrison. 1986. Immunologic activity of lipopolysaccharides released from macrophages after the uptake
of intact E. coli in vitro. J. Immunol. 136:2924.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
Acknowledgments
We thank J. Ewbank, I. Moryion, and S. Méresse for critically reading the
manuscript.
29.
References
30.
1. Rietschel, E. T., T. Kirikae, F. U. Schade, U. Mamat, G. Schmidt, H. Loppnow,
A. J. Ulmer, U. Zahringer, U. Seydel, and F. Di Padova. 1994. Bacterial endotoxin: molecular relationships of structure to activity and function. FASEB J.
8:217.
2. Russel, R. R. B. 1977. Free endotoxin. Microbios Lett. 2:125.
3. Fox, E. S., P. Thomas, and S. A. Broitman. 1987. Comparative studies of endotoxin uptake by isolated rat Kupffer and peritoneal cells. Infect. Immun. 55:2962.
4. Hampton, R. Y., D. T. Golenbock, M. Penman, M. Krieger, and C. R. Raetz.
1991. Recognition and plasma clearance of endotoxin by scavenger receptors.
Nature 352:342.
5. Poussin, C., M. Foti, J. L. Carpentier, and J. Pugin. 1998. CD14-dependent endotoxin internalization via a macropinocytic pathway. J. Biol. Chem. 273:20285.
6. Kang, Y. H., R. S. Dwivedi, and C. H. Lee. 1990. Ultrastructural and immunocytochemical study of the uptake and distribution of bacterial lipopolysaccharide
in human monocytes. J. Leukocyte Biol. 48:316.
7. Kang, Y. H., M. Carl, R. K. Maheshwari, L. P. Watson, L. Yaffe, and
P. M. Grimley. 1988. Incorporation of bacterial lipopolysaccharide by human
Leu-11a1 natural killer cells: ultrastructural and functional correlations. Lab.
Invest. 58:196.
8. Garcia-del Portillo, F., M. A. Stein, and B. B. Finlay. 1997. Release of lipopolysaccharide from intracellular compartments containing Salmonella typhimurium
to vesicles of the host epithelial cell. Infect. Immun. 65:24.
9. Duncan, R. L. Jr., and D. C. Morrison. 1984. The fate of E. coli lipopolysaccharide after the uptake of E. coli by murine macrophages in vitro. J. Immunol.
132:1416.
10. Risco, C., J. L. Carrascosa, and M. A. Bosch. 1991. Uptake and subcellular
distribution of Escherichia coli lipopolysaccharide by isolated rat type II pneumocytes. J. Histochem. Cytochem. 39:607.
11. Kriegsmann, J., S. Gay, and R. Brauer. 1993. Endocytosis of lipopolysaccharide
in mouse macrophages. Cell. Mol. Biol. 39:791.
12. Odeyale, C. O., and Y. H. Kang. 1988. Biotinylation of bacterial lipopolysaccharide and its applications to electron microscopy. J. Histochem. Cytochem.
36:1131.
13. Wuorela, M., S. Jalkanen, P. Toivanen, and K. Granfors. 1993. Yersinia lipopolysaccharide is modified by human monocytes. Infect. Immun. 61:5261.
14. Lang, T., M. T. Tassin, and A. Ryter. 1988. Bacterial antigen immunolabeling in
macrophages after phagocytosis and degradation of Bacillus subtilis. Infect. Immun. 56:468.
15. Dacosta, B., A. Ryter, J. Mounier, and P. Sansonetti. 1990. Immunodetection of
lipopolysaccharide in macrophages during the processing of noninvasive Shigella
dysenteriae. Biol. Cell 69:171.
16. Leyva-Cobian, F., I. M. Outschoorn, E. Carrasco-Marin, and
C. Alvarez-Dominguez. 1997. The consequences of the intracellular retention of
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
45.
46.
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
6790
The Journal of Immunology
63. Munford, R. S., C. L. Hall, and P. D. Rick. 1980. Size heterogeneity of Salmonella typhimurium lipopolysaccharides in outer membranes and culture supernatant membrane fragments. J. Bacteriol. 144:630.
64. Freudenberg, M. A., A. Fomsgaard, I. Mitov, and C. Galanos. 1989. ELISA for
antibodies to lipid A, lipopolysaccharides and other hydrophobic antigens. Infection 17:322.
65. Hitchcock, P. J., and T. M. Brown. 1983. Morphological heterogeneity among
Salmonella lipopolysaccharide chemotypes in silver-stained polyacrylamide gels.
J. Bacteriol. 154:269.
66. Caroff, M., D. R. Bundle, M. B. Perry, J. W. Cherwonogrodzky, and
J. R. Duncan. 1984. Antigenic S-type lipopolysaccharide of Brucella abortus
1119-3. Infect. Immun. 46:384.
67. Granfors, K., S. Jalkanen, P. Toivanen, J. Koski, and A. A. Lindberg. 1992.
Bacterial lipopolysaccharide in synovial fluid cells in Shigella triggered reactive
arthritis. J. Rheumatol. 19:500.
68. Granfors, K. 1992. Do bacterial antigens cause reactive arthritis? Rheum. Dis.
Clin. North Am. 18:37.
69. Granfors, K., S. Jalkanen, A. A. Lindberg, O. Maki-Ikola, R. von Essen,
R. Lahesmaa-Rantala, H. Isomaki, R. Saario, W. J. Arnold, and A. Toivanen.
1990. Salmonella lipopolysaccharide in synovial cells from patients with reactive
arthritis. Lancet 335:685.
70. Beckman, E. M., S. A. Porcelli, C. T. Morita, S. M. Behar, S. T. Furlong, and
M. B. Brenner. 1994. Recognition of a lipid antigen by CD1-restricted ab1 T
cells. Nature 372:691.
71. Sieling, P. A., D. Chatterjee, S. A. Porcelli, T. I. Prigozy, R. J. Mazzaccaro,
T. Soriano, B. R. Bloom, M. B. Brenner, M. Kronenberg, and P. J. Brennan.
1995. CD1-restricted T cell recognition of microbial lipoglycan antigens. Science
269:227.
72. Spada, F. M., Y. Koezuka, and S. A. Porcelli. 1998. CD1d-restricted recognition
of synthetic glycolipid antigens by human natural killer T cells. J. Exp. Med.
188:1529.
73. Burdin, N., L. Brossay, Y. Koezuka, S. T. Smiley, M. J. Grusby, M. Gui,
M. Taniguchi, K. Hayakawa, and M. Kronenberg. 1998. Selective ability of
mouse CD1 to present glycolipids: a-galactosylceramide specifically stimulates
Va141 NK T lymphocytes. J. Immunol. 161:3271.
74. Kawano, T., J. Cui, Y. Koezuka, I. Toura, Y. Kaneto, K. Motoki, H. Ueno,
R. Nakagawa, H. Sato, E. Kondo, H. Koseki, and M. Taniguchi. 1997. CD1drestricted and TCR-mediated activation of Va14 NKT cells by glycosylceramides. Science 278:1626.
75. Mattern, T., A. Thanhäuser, N. Reiling, K.-M. Toellner, M. Duchrow,
S. Kusumoto, E. T. Rietschel, M. Enrst, H. Brade, H.-D. Flad, and A. J. Ulmer.
1994. Endotoxin and lipid A stimulate proliferation of human T cells in the
presence of autologous monocytes. J. Immunol. 153:2996.
76. Tough, D. F., S. Sun, and J. Sprent. 1997. T cell stimulation in vivo by lipopolysaccharide (LPS). J. Exp. Med. 185:2089.
77. Mattern, T., H.-D. Flad, L. Brade, E. T. Rietschel, and A. J. Ulmer. 1998. Stimulation of human T lymphocytes by LPS is MHC unrestricted, but strongly dependent on B7 interactions. J. Immunol. 160:3412.
78. Castro, A., V. Bemer, A. Nobrega, A. Coutinho, and P. Truffa-Bachi. 1998.
Administration to mouse of endotoxin from Gram-negative bacteria leads to activation and apoptosis of T lymphocytes. Eur. J. Immunol. 28:488.
Downloaded from http://www.jimmunol.org/ by guest on June 14, 2017
47. Montgomery, R. R., P. Webster, and I. Mellman. 1991. Accumulation of indigestible substances reduces fusion competence of macrophage lysosomes. J. Immunol. 147:3087.
48. Wurfel, M. M., B. G. Monks, R. R. Ingalls, R. L. Dedrick, R. Delude, D. Zhou,
N. Lamping, R. R. Schumann, R. Thieringer, M. J. Fenton, et al. 1997. Targeted
deletion of the lipopolysaccharide (LPS)-binding protein gene leads to profound
suppression of LPS responses ex vivo, whereas in vivo responses remain intact.
J. Exp. Med. 186:2051.
49. Wang, P. Y., R. L. Kitchens, and R. S. Munford. 1995. Bacterial lipopolysaccharide binds to CD14 in low-density domains of the monocyte-macrophage
plasma membrane. J. Inflamm. 47:126.
50. Morrison, D. C., and J. A. Rudbach. 1981. Endotoxin-cell-membrane interactions
leading to transmembrane signaling. Contemp. Top. Mol. Immunol. 8:187.
51. Price, R. M., and D. M. Jacobs. 1986. Fluorescent detection of lipopolysaccharide
interactions with model membranes. Biochim. Biophys. Acta 859:26.
52. Schromm, A. B., K. Brandenburg, E. T. Rietschel, H. D. Flad, S. F. Carroll, and
U. Seydel. 1996. Lipopolysaccharide-binding protein mediates CD14-independent intercalation of lipopolysaccharide into phospholipid membranes. FEBS
Lett. 399:267.
53. Schromm, A. B., K. Brandenburg, H. Loppnow, U. Zahringer, E. T. Rietschel,
S. F. Carroll, M. H. Koch, S. Kusumoto, and U. Seydel. 1998. The charge of
endotoxin molecules influences their conformation and IL-6-inducing capacity.
J. Immunol. 161:5464.
54. Korn, A., Z. Rajabi, B. Wassum, W. Ruiner, and K. Nixdorff. 1995. Enhancement
of uptake of lipopolysaccharide in macrophages by the major outer membrane
protein OmpA of Gram-negative bacteria. Infect. Immun. 63:2697.
55. Pagano, R. E., O. C. Martin, H. C. Kang, and R. P. Haugland. 1991. A novel
fluorescent ceramide analogue for studying membrane traffic in animal cells:
accumulation at the Golgi apparatus results in altered spectral properties of the
sphingolipid precursor. J. Cell Biol. 113:1267.
56. Koval, M., and R. E. Pagano. 1990. Sorting of an internalized plasma membrane
lipid between recycling and degradative pathways in normal and Niemann-Pick,
type A fibroblasts. J. Cell Biol. 111:429.
57. Koval, M., and R. E. Pagano. 1989. Lipid recycling between the plasma membrane and intracellular compartments: transport and metabolism of fluorescent
sphingomyelin analogues in cultured fibroblasts. J. Cell Biol. 108:2169.
58. Joseph, C. K., S. D. Wright, W. G. Bornmann, J. T. Randolph, E. R. Kumar,
R. Bittman, J. Liu, and R. N. Kolesnick. 1994. Bacterial lipopolysaccharide has
structural similarity to ceramide and stimulates ceramide-activated protein kinase
in myeloid cells. J. Biol. Chem. 269:17606.
59. Thieblemont, N., and S. D. Wright. 1997. Mice genetically hyporesponsive to
lipopolysaccharide (LPS) exhibit a defect in endocytic uptake of LPS and ceramide. J. Exp. Med. 185:2095.
60. Mellman, I., P. Pierre, and S. Amigorena. 1995. Lonely MHC molecules seeking
immunogenic peptides for meaningful relationships. Curr. Opin. Cell Biol. 7:564.
61. Escola, J. M., E. Moreno, P. Chavrier, and J. P. Gorvel. 1994. The O-chain of
Brucella abortus lipopolysaccharide induces SDS-resistant MHC class II molecules in mouse B cells. Biochem. Biophys. Res. Commun. 203:1230.
62. Rietschel, E. T., U. Seydel, U. Zahringer, U. F. Schade, L. Brade, H. Loppnow,
W. Feist, M. H. Wang, A. J. Ulmer, and H. D. Flad. 1991. Bacterial endotoxin:
molecular relationships between structure and activity. Infect. Dis. Clin. North
Am. 5:753.
6791