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Molecular Host Defense Laboratory, Departments of
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Internal Medicine (Infectious Disease Division) and
Microbiology, University of Texas Southwestern Medical Center, Dallas, TX 75235
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The mechanism of internalization of mCD14-bound LPS has not been described. It may enter cells via clathrin-coated pits (11, 12) or possibly via the noncoated invaginations reported to internalize other glycosylphosphatidylinositol-anchored proteins (13, 14, 15, 16). LPS then moves into acidic intracellular compartments where the lipid A moiety may be partially degraded in ways that modify its biologic activity (17). LPS has been shown to move from the plasma membrane to intracellular vesicles (4) that, in macrophages, may reach a perinuclear location (18). There is also evidence that LPS may remain associated with the plasma membrane for extended periods of time (19) and that cells may release bioactive LPS (20).
The biologic importance of LPS internalization and its relationship to signaling are disputed. Much experimental evidence suggests that internalized LPS undergoes enzymatic degradation or physical sequestration (17), processes that should reduce its signal potency (5). LPS-induced signals may regulate these processes. For example, LPS can down-regulate its own dephosphorylation by macrophages (21), and according to one recent report (18), C3H/HeJ macrophages, which fail to respond to LPS, show alterations in the movement of intracellular LPS-containing vesicles. While some evidence suggests that LPS must be present in intracellular vesicles before certain signals are transduced (6, 18, 22), other data argue that the LPS that undergoes internalization by monocytes is not involved in signaling (5). Clarification of these issues will require knowledge of the mechanisms by which LPS is internalized and transported within the cell.
In this report we show that the kinetics with which mCD14-bound LPS is internalized are influenced prominently by the initial LPS aggregation state, but not by cellular responses to the LPS.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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THP-1 cells (23) were obtained from D. Altieri (Scripps Research Institute, La Jolla, CA) and cultured as previously described (8). To induce CD14 expression, the cells were either differentiated by culture in 0.05 µM 1,25-dihydroxyvitamin D3 for 48 to 96 h (7, 8) or stably transfected by electroporation with human CD14 cDNA (a gift from D. Golenbock, Boston University, Boston, MA) cloned into pRc/RSV (Invitrogen, San Diego, CA). Bulk populations of stably transformed cells were selected in 0.5 mg/ml G418, and cells expressing CD14 were isolated using a fluorescence-activated cell sorter (FACStar Plus, Becton Dickinson Immunocytometry, San Jose, CA) and expanded in culture. Thioglycolate-elicited peritoneal macrophages were isolated as previously described (24) from C3H/HeJ and C3H/HeN mice (The Jackson Laboratory, Bar Harbor, ME). CHO cells stably transfected with recombinant human LPS binding protein (rLBP) or empty vector (pRc/RSV) were provided by P. Tobias (Scripps Research Institute). The cells (CHO-rLBP or CHO-RSV) were cultured in serum-free medium (CHO-S-SFM II, Life Technologies, Grand Island, NY).
Reagents
CHO-rLBP and CHO-RSV culture supernatants were tested for their ability to promote [3H]LPS (100 ng/ml) binding to CD14 on THP-1 cells, and a dilution of the CHO-rLBP supernatant that rapidly enhanced LPS binding was chosen for subsequent experiments. CHO-RSV supernatant had no LPS transfer activity. Purified recombinant human soluble CD141–356 (sCD14) was a gift from R. Thieringer (Merck, Rahway, NJ). Anti-CD14 mAb 26ic (IgG2b) was provided by D. Golenbock (Boston University). FITC-conjugated goat anti-mouse IgG (H+L) F(ab')2 was obtained from Tago (Burlingame, CA). RPMI 1640, Cellgro Complete serum-free medium, and G418 were purchased from Mediatech (Herndon, VA). Proteinase K (from Tritirachium album), cell culture-tested BSA, nonimmune IgG2b control Ab MOPC-141, PMSF, and 1,4-diazabicyclo-(2, 2, 2)octane were obtained from Sigma Chemical Co. (St. Louis, MO). Phosphatidylinositol-specific phospholipase C (PI-PLC) from Bacillus cereus was purchased from Boehringer Mannheim (Indianapolis, IN).
LPS preparations
Escherichia coli LCD25 [3H]LPS (1.5 x 106 dpm/µg) was biosynthetically labeled and isolated as previously described (25). For derivatization, unlabeled LCD25 LPS was obtained from List Biological Laboratories (Campbell, CA) and repurified (26) to remove trace protein contamination. After repurification, contaminating protein could not be detected on silver-stained SDS-PAGE gels after loading 10 µg of LPS/lane. The LPS was fluoresceinated with FITC (Molecular Probes, Eugene, OR) (27) as follows. One hundred micrograms of LPS containing tracer amounts of [3H]LPS were mixed with 0.1 M borate buffer, pH 10.5, containing 0.3% sodium deoxycholate (Sigma Ultrapure), 1 mM EDTA, and 10 mg/ml FITC, and the mixture was incubated in the dark for 3 h at 37°C. Unbound FITC was removed by extensive dialysis against 0.9% NaCl with 10 mM Tris (Cl), pH 7.5, at 4°C. The molar ratio of FITC to LPS was 0.36.
Preparation of monomeric and aggregated LPS
Aggregated LPS (Ag-LPS) was prepared by diluting [3H]LPS stock suspensions to a concentration of 10 to 20 µg/ml in RPMI-HB and dispersing with a probe sonicator for 1 to 2 s (model 450, Branson Ultrasonics, Danbury, CT). Partially disaggregated LPS (DAg-LPS) was made in the same way, except that the RPMI-HB was replaced by HNEB. Monomeric LPS-sCD14 complexes (28) were prepared by mixing DAg-LPS with a 25-fold excess (by weight) of sCD14 (usually 300–500 µg sCD14/ml, final concentration) and incubating overnight at 37°C. For some experiments, LPS-sCD14 complexes were made in the presence of LBP by mixing LPS and sCD14 as described above with an equal volume of CHO-rLBP supernatant for 10 min at 37°C. LPS-sCD14 complexes generated by either method were filtered through a 100-kDa cut-off membrane (Microcon-100, Amicon, Beverly, MA) to minimize the presence of aggregates.
Sucrose gradient analysis of LPS
Sucrose gradients were made as previously described (29). Samples (0.2 ml) containing [3H]LPS were loaded on top of 4.7-ml gradients and centrifuged at 150,000 x g for 4.5 h at room temperature. Fractions (0.35 ml) were collected from the bottom with an 18-gauge needle, and 3H disintegrations per minute were measured in the presence of SDS and EDTA as previously described (7).
Cell stimulation assays
THP-1 cells were washed three times with cold RPMI 1640 and
resuspended in SFM (Cellgro Complete serum-free medium containing 20 mM
HEPES buffer, pH 7.4, and 0.3 mg/ml BSA) at 5 to 7 x
106 cells/ml. LPS was diluted into CHO-rLBP or CHO-RSV
supernatant and incubated for 5 min at 37°C before addition to the
cells. The cells (90 µl) were warmed to 37°C for 5 min, 10 µl of
LPS were added, and the incubation was continued at 37°C for 1 or
2 h. The cells were chilled on ice, the suspensions were
centrifuged, and the culture supernatants were assayed for IL-8 (DuoSet
ELISA Development System, Genzyme, Cambridge, MA). NF-
B was measured
by gel-shift assay in nuclear extracts as previously described (8).
Relative amounts of radioactivity in the gel-shifted bands were
measured using a PhosphorImager SF (Molecular Dynamics, Sunnyvale, CA)
and expressed in arbitrary units.
LPS internalization assays
Cell binding. DAg-LPS ([3H]LPS or FITC-LPS) was used unless otherwise stated. The LPS in HNEB was diluted 1/10 with CHO-RSV or CHO-rLBP supernatant and incubated for 5 min at 37°C immediately before addition to the cells. LPS-sCD14 complexes were diluted in PBS with 0.3 mg/ml BSA. The cells were harvested in suspension, washed three times with cold RPMI 1640, and resuspended in SFM at 5 to 7 x 106 cells/ml. Cells (90 µl) were warmed to 37°C, 10 µl of LPS were added, and the mixtures were incubated at 37°C for 1 to 60 min. For binding LPS to cells on ice, the DAg-LPS-CHO LBP supernatant mixtures were incubated for 5 min at 37°C, chilled on ice, mixed with the cells, and allowed to bind for 15 min on ice. The cells were then washed, resuspended in 100 µl of SFM, and warmed to 37°C for various times. Internalization was stopped by adding 1 ml of ice-cold PBS. The cells were pelleted by centrifugation (750 x g for 2 min at 0–4°C) and washed with 1 ml of cold PBS. Adherent macrophages were isolated in 24-well culture plates (Costar, Cambridge, MA), washed three times with RPMI 1640, and warmed to 37°C for 5 min in 270 µl of SFM. [3H]LPS in CHO-LBP or CHO-RSV supernatant (30 µl) was added to the cells, and the incubation was continued at 37°C for various times. The cells were then placed on ice, aspirated, and washed twice with 1 ml of ice-cold PBS. Surface-exposed and internal LPS were measured as described below.
Protease protection assay. The washed cells were resuspended in 1 ml of ice-cold 0.02% proteinase K and kept on ice for 30 min. The cells were pelleted by centrifugation, and 300 µl of supernatant were removed for 3H counting. The cells were resuspended in 200 µl of PBS containing 0.5 mM PMSF, and 100 µl of cells were counted. The protein content of each cell suspension was measured using Bio-Rad protein assay reagent (Bio-Rad, Hercules, CA). Proteinase K treatment did not increase cell permeability to trypan blue or decrease cell number (not shown), although some of the cells that were already permeable to trypan blue were digested by the protease. For removal of surface-exposed LPS with PI-PLC instead of proteinase K, the washed cells were resuspended in 100 µl of HNE buffer (20 mM HEPES (pH 7.4), 150 mM NaCl, and 1 mM EDTA) containing 0.5 U of PI-PLC. The mixture was incubated for 1 h on ice, 900 µl of cold PBS were added, and the cells were pelleted by centrifugation. Cell-associated and released [3H]LPS were measured as described above. For adherent murine macrophages, the washed cells were incubated on ice with 500 µl of 0.02% proteinase K in PBS. The protease solution was saved, and the cells were resuspended in 200 µl of PBS using a rubber policeman. [3H]LPS and cell protein content were measured as described above.
Fluorescence quenching assay. After incubating cells with FITC-LPS in suspension, the cells were washed as described above, resuspended in 100 µl of SFM, and split into 50-µl aliquots. Rabbit anti-fluorescein (Texas Red conjugate; Molecular Probes, Eugene, OR) was added to one aliquot (50 µg/ml, final concentration), and the mixtures were incubated on ice for 30 min. The cells were then washed with cold PBS. To prevent quenching of fluorescein, the intracellular pH was raised by fixing the cells in 1 ml of 3% paraformaldehyde in 100 mM sodium phosphate, pH 7.4, for 30 min on ice, centrifuging, and resuspending the cells in cold PBS containing 20 mM Tris, pH 8.0. The mean fluorescence intensity (MFI) of FITC-LPS in each cell population was measured by a flow cytometer (FACScan, Becton Dickinson Immunocytometry, San Jose, CA). We found the same relationship between green (530 nm) and orange (585 nm) fluorescence emissions in cell populations that had internalized FITC-LPS (not shown) as in those that had not internalized FITC-LPS, providing evidence that intracellular fluorescein was not quenched. In some experiments, anti-fluorescein was replaced by 1 ml of 0.02% proteinase K in PBS to remove surface-exposed FITC-LPS.
Measurement of mCD14 by FACS analysis
THP-1 cells were washed twice with cold PBS containing 0.5 mM PMSF and resuspended in 90 µl of blocking buffer (PBS with 0.3 mg/ml BSA, 10% heat-inactivated normal goat serum, and 0.5 mM PMSF) for 10 min on ice. The mCD14 was stained with anti-CD14 mAb 26ic or control IgG2b (10 µg/ml) for 45 min on ice followed by FITC-conjugated goat anti-mouse IgG F(ab')2 (1/200) for 45 min. The MFI in each cell population was measured by FACS analysis.
Laser confocal microscope imaging
Cells were incubated with 200 ng/ml FITC-LPS prepared in CHO-LBP or CHO-RSV supernatant for various times at 37°C in SFM as described above. The cells were washed twice with cold PBS and resuspended in 200 µl of 4% paraformaldehyde in 100 mM sodium phosphate, pH 7.4, for 30 min on ice. The cells were then centrifuged onto poly-L-lysine-coated slides for 5 min at room temperature and washed in cold PBS. The slides were rinsed in distilled water, aspirated dry, and mounted with one drop of mounting solution (9 vol of glycerol, 1 vol of 1 M Tris (Cl), pH 8.6, and 2.5% 1,4-diazabicyclo-(2, 2, 2)octane) with a 22- x 22-mm glass coverslip (no. 1, Fisher Scientific, Pittsburgh, PA). The cells were viewed with an MRC-1024 laser confocal imaging system (Bio-Rad). Sequential optical sections (1 µm) were observed, and the resolution of the digital images was 512 x 512 pixels (1 pixel = 0.155 µm).
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Binding and internalization measurements in this study were
restricted to LPS that binds initially to mCD14 in the presence of
either LBP or sCD14. We (7) and others (30) have previously shown that
LBP promotes LPS binding to mCD14 in THP-1 cells. Hailman et al. (28)
showed that [3H]LPS-sCD14 complexes can rapidly
transfer LPS to unoccupied sCD14 molecules in the absence of LBP and
further showed indirect evidence that sCD14 transfers LPS to mCD14.
Using CD14-transfected THP-1 cells, we found that [3H]LPS
from [3H]LPS-sCD14 complexes binds to mCD14 in the
absence of LBP (Fig. 1
). In contrast,
mCD14-negative cells that were transfected with empty vector did not
measurably bind LPS from LPS-sCD14 complexes. The LPS bound to mCD14
saturably and with high apparent affinity. The apparent equilibrium
dissociation constant (Kd) of 31 ±
3 ng/ml (n = 2) or 7.8 nM was very similar to
that obtained for [3H]LPS-LBP complexes
(Kd = 33 ng/ml or 8.2 nM) (Ref. 7 and
data not shown). The actual Kd (not determined)
may differ from the apparent Kd reported here,
since the affinity of the LPS-sCD14 interaction and the equilibrium
between LPS-mCD14 and LPS-sCD14 were not taken into account. The
binding capacity (6.2 x 105 molecules of LPS/cell) is
similar to the number of mCD14 molecules expressed per cell (Ref. 7 and
data not shown).
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The protease protection assay is based on the ability of
proteinase K to remove surface-exposed LPS at the temperature of ice
while leaving the plasma membrane intact. As shown in Table I, proteinase K removed nearly all the
[3H]LPS that bound to the cells whether they had
been incubated with [3H]LPS-LBP or with
[3H]LPS-sCD14 complexes. Likewise, proteinase K removed
virtually all the mCD14 from the cells. Treatment of the cells with
PI-PLC, which cleaves CD14 from its glycosylphosphatidylinositol
anchor, removed less [3H]LPS than treatment with
proteinase K. However, PI-PLC also removed significantly less mCD14
from the cells; this may account for its lower efficiency at removing
[3H]LPS. As shown in Table I, the percentages of the
total mCD14 (73%) and [3H]LPS (derived from sCD14
complexes; 69%) that were removed by PI-PLC were virtually the same.
PI-PLC removed significantly less [3H]LPS (from LBP
complexes) than mCD14, however, suggesting that some of the
LBP-delivered LPS may interact with molecule(s) other than mCD14 on the
cell surface. Although the [3H]LPS was labeled entirely
in its fatty acyl chains (25), deacylation of the cell-associated LPS
occurred very slowly in these cells (<1.5% in 1 h; data not
shown). This minor loss of 3H from the
[3H]LPS should not significantly affect the measurements
described here.
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shows
representative fluorescence histograms from an experiment in which
THP-1 cells were exposed to FITC-LPS for 10 min at 37°C. MFI data
were corrected (
MFI) for the autofluorescence of the cells
(a; MFI = 4.6). LPS binding in the absence of
LBP (b; MFI = 1.2) was only slightly higher
than the autofluorescence of the cells. After incubation with FITC-LPS
in the presence of LBP, internalized LPS was measured by exposing the
cells to the anti-fluorescein Ab (d; MFI
= 13.8). Surface-exposed LPS was determined by measuring the total
cell-associated LPS (c; MFI = 29.3) and
subtracting internal LPS (c - d;
29.3 - 13.8 = 15.5). At least 90% of the FITC-LPS that
bound to metabolically inhibited cells was quenched by the Ab (not
shown).
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Figure 3 shows LPS binding and
internalization measured by protease protection
(A) and by fluorescence quenching
(B) in cells that were exposed to LPS for the
indicated times at 37°C. MFI data for FITC-LPS are shown on a linear
scale. LPS bound rapidly to mCD14 (maximal in 2 min) under these
conditions. The kinetics of internalization were biphasic. Most of the
LPS internalization occurred rapidly (within 5 min), followed by
markedly slower internalization with very little additional
accumulation of LPS inside the cells. Similar results were obtained for
FITC-LPS internalization by freshly isolated human monocytes (not
shown).
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shows that after exposing THP-1 cells
to FITC-LPS for 20 min at 37°C, protease treatment and fluorescence
quenching gave virtually identical estimates of surface-exposed
FITC-LPS. The results indicate that the LPS was not measurably
transferred from mCD14 to a protease-resistant site. They also confirm
that proteinase K treatment does not remove internal or sequestered LPS
that is inaccessible to the gentler Ab quenching method.
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shows the proportions of the total
cell-associated [3H]LPS that are internal (protease
resistant) or on the cell surface (protease sensitive) at each time
point. Internalization was rapid during the first 5 min of rewarming
and slowed thereafter. The biphasic internalization kinetics were
similar to those observed in Figure 3
. Rewarming the cells for 20 min
resulted in <10% release of surface-bound LPS into the medium (not
shown). Similar results were obtained whether CD14-transfected cells
(Fig. 4) or differentiated THP-1 cells (Fig. 8) were used. We found
that efficient binding of LPS to cells on ice required binding the LPS
to LBP at 37°C before chilling on ice (5). Binding under these
conditions was nearly maximal in 15 min (not shown).
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We used laser confocal microscopy to visualize surface and
internal FITC-LPS after exposure to THP-1 cells for various times at
37°C (Fig. 5). After 1 min, virtually
all the LPS was diffusely distributed on the plasma membrane (Fig. 5A). Treatment of the cells with proteinase K almost
completely removed the FITC-LPS, although a few focal accumulations
could be found (Fig. 5E). These foci became more
numerous and larger with increasing incubation times and were observed
in proteinase K-treated cells, suggesting that they represent
internalized LPS. We analyzed a series of 10 optical sections of each
sample at 1-µm intervals and confirmed the intracellular location of
many of the fluorescent foci (not shown). The resolution of this
method, however, is not sufficient to distinguish whether the foci at
the cell periphery are on the outer or the inner surface of the plasma
membrane, and we cannot exclude the possibility that some of these foci
represent LPS that accumulates in membrane invaginations. Most of the
internalized (protease-resistant) LPS was found near the cell surface
at early time points (1–5 min; A and B,and E and F), but at later time
points (20–45 min; C and D, and G and
H), much of the LPS was clearly separated from the
plasma membrane. Although we cannot quantitate LPS internalization
kinetics by this method, the confocal images confirm that FITC-LPS is
internalized by the cells and that much of the LPS becomes well
separated from the plasma membrane.
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[3H]LPS was prepared in various aggregation
states and subjected to sucrose density gradient analysis (Fig. 6). LPS stock suspensions that were
diluted directly into Ca2+- and Mg2+-containing
medium (RPMI 1640) remained highly aggregated even after probe
sonication and exposure to LBP, and this LPS (Ag-LPS) migrated through
the gradient to the bottom of the centrifuge tube (fraction 0).
Monomeric LPS, prepared by incubating [3H]LPS overnight
with an excess of purified soluble CD14 (28) in the presence or the
absence of LBP, remained at the top of the sucrose gradient. DAg-LPS
(used in the experiments above), prepared by diluting and sonicating
stock suspensions in the presence of EDTA, moved slightly farther down
the gradients than monomeric LPS, suggesting that the molecules were
somewhat aggregated. When DAg-LPS was diluted in Ca2+- and
Mg2+-containing medium, the aggregation state increased to
a variable extent but did not revert to the highly aggregated state of
Ag-LPS. Exposure to LBP under these conditions did not disaggregate the
LPS to the extent observed with LPS-sCD14 complexes (not shown).
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Monomeric LPS bound to mCD14 in the presence (not shown) or
the absence (Fig. 7A)
of LBP at a rate similar to that at which DAg-LPS bound in the presence
of LBP (Fig. 7B; maximal in 1–2 min). While nearly half the
maximal amount of Ag-LPS bound to the cells in 1 to 2 min, its binding
reached a maximum more slowly (10–30 min). In contrast, the cells
internalized aggregated LPS extremely rapidly (70–80% of the total
LPS that bound to the cells was internalized in 1 min; Fig. 7C), whereas LPS monomers were internalized very
slowly (Fig. 7A). Partially disaggregated LPS was
internalized at an intermediate rate (Fig. 7B).
Although the initial rate of monomeric LPS internalization was markedly
slower than that of partially disaggregated LPS, the second phase
kinetics were similar (Fig. 7D). Compared with the
LPS monomers, approximately sixfold more LPS from the aggregated
preparation was internalized during the first 20 min of incubation
(Fig. 7, A and C).
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B), but they did not exhibit the dramatic
differences seen with aggregated and disaggregated
[3H]LPS preparations. This may be explained by the fact
that the derivatization procedure for FITC-LPS involves disaggregating
the LPS with deoxycholate. LPS treated in this way may not completely
reaggregate to its original state. THP-1 cells internalized much less
FITC-LPS from monomeric FITC-LPS-sCD14 complexes (not shown), in
keeping with the results of the protease protection assay for
[3H]LPS (Fig. 7A). Aggregation state has little effect on LPS stimulatory potency
The stimulatory potencies of the LPS preparations were assessed by
their abilities to induce IL-8 production and NF-B translocation in
differentiated THP-1 cells. When incubated with cells at the submaximal
stimulatory concentration of 1 ng/ml, the three preparations exhibited
similar responses (IL-8 production and NF-B translocation; Table III). The experiments were repeated with
similar results.
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Having previously shown that a 3-h pre-exposure to LPS
desensitized THP-1 cells to restimulation of p42 protein tyrosine
phosphorylation by LPS (31), we tested whether LPS internalization
kinetics could be altered by any of the stimulatory or desensitizing
effects of LPS pre-exposure. THP-1 cells were incubated with unlabeled
LPS in the presence of LBP at 37°C before adding
[3H]LPS plus LBP or monomeric
[3H]LPS-sCD14 complexes. The rate and the extent of
internalization of LPS aggregates (Fig. 8A) or LPS monomers
(not shown) were not significantly altered by pre-exposure to 2 or 100
ng/ml LPS for 3 h, although LPS binding was slightly decreased in
some experiments (not shown). Similar results were obtained after short
(5-min) pre-exposure to 50 ng/ml LPS (not shown). We next asked whether
an LPS-specific inhibitor would alter LPS internalization. We
previously showed that the tetra-acyl lipid A analogue, LA-14-PP, can
inhibit LPS responses without inhibiting LPS uptake by mCD14 (8). As
shown in Figure 8B, a concentration of LA-14-PP that
strongly inhibited LPS-induced IL-8 production did not alter
[3H]LPS internalization. IL-8 production measured at
2 h was 1.55 ± 0.12 ng/ml (no LPS), 6.49 ± 0.19 (LPS),
2.78 ± 0.17 (LA-14-PP plus LPS), and 1.78 ± 0.09 (LA-14-PP
alone). The experiment was repeated with similar results. LA-14-PP also
did not inhibit the internalization of monomeric LPS from
[3H]LPS-sCD14 complexes (not shown).
Finally, we measured internalization by macrophages from LPS-responsive
(C3H/HeN) and LPS-hyporesponsive (C3H/HeJ) mice. Figure 9 shows that the macrophages internalized
LPS with identical kinetics whether they were derived from HeN or HeJ
mice, suggesting that LPS-induced responses do not alter the ability of
cells to internalize LPS. The LPS hyporesponsiveness of the HeJ
macrophages was confirmed by stimulating the HeN and HeJ macrophages
for 4 h with 10 ng/ml LPS and measuring IL-6 production by ELISA.
Using cells from the same experiment (Fig. 9), the HeN macrophages
released 67 ± 3 ng/ml IL-6 into the culture supernatant, whereas
the HeJ macrophages released <7 ng/ml. Taken together, the data in
Figures 8 and 9 suggest that mCD14-bound LPS is internalized by a
constitutive cellular process that is not affected by LPS-induced
responses or desensitization.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Our initial goal was therefore to develop and test two independent, quantitative methods for measuring LPS internalization. We found that the protease protection and fluorescence quenching assays gave remarkably similar results, and laser confocal microscope images confirmed that the cells had internalized the LPS. These methods should be useful for quantitating LPS internalization in a variety of cell types.
It is important to note that the term internalization, as used here,
refers to the movement of surface-exposed LPS to sites that are
inaccessible to proteinase K or anti-fluorescein. The kinetics of this
process may be quite different from the kinetics of subsequent
processes that move LPS to various intracellular compartments. For
example, Ward et al. (37, 38) showed that different ligands were
internalized by their respective receptors on macrophages at vastly
different rates, yet the kinetics of intracellular traffic were the
same for each ligand. In neutrophils, LPS was shown to move slowly to
an endocytic compartment that comigrated on Percoll gradients with
vesicles that contain acyloxyacyl hydrolase, an enzyme that partially
degrades LPS (4). The process by which internalized LPS moves into this
compartment, however, might have very different kinetics from the
process by which LPS enters the cell. Likewise, the recently reported
(18) defect in intracellular LPS traffic in C3H/HeJ macrophages appears
to be distal to LPS internalization. As we show in Figure 9, LPS
internalization occurs with identical kinetics in C3H/HeN and C3H/HeJ
macrophages.
The aggregation state had a dramatic influence on the initial rate and
extent of CD14-dependent internalization of this amphipathic ligand.
Highly aggregated LPS was internalized extremely rapidly, whereas
monomeric LPS was internalized slowly. DAg-LPS was internalized at an
intermediate rate. The initial phase of internalization of large LPS
aggregates (70% of the cell-associated LPS within 1 min) occurred at a
rate similar to that reported for the internalization of polyvalent
mannosylated ligands by the mannose receptor on sinusoidal endothelial
cells (t1/2 = 10 s) (39) and
on rabbit alveolar macrophages
(t1/2 = 1 min) (37). Such rapid
internalization could be promoted by the ability of each ligand
aggregate to cluster many receptor molecules, which may result in
binding to endocytic structures with greater avidity. We cannot rule
out the possibility, however, that the rapid sequestration of LPS
aggregates from the actions of proteinase K or anti-fluorescein may
be due to the inability of these proteins to access some
ligand-receptor aggregates that are still surface exposed. Evidence
that the assays measure LPS internalization includes the observations
that 1) rapid internalization of LPS aggregates is energy dependent
(Table I and data not shown), whereas LPS aggregation and binding to
CD14 are not; 2) proteinase K and anti-fluorescein act by very
different mechanisms (removal of LPS by degradation of mCD14 and
possibly of LBP in contrast to quenching of exposed fluorescein groups
on the LPS), but yield identical measurements of surface-exposed LPS;
and 3) as mentioned above, the rate of internalization of aggregated
LPS is similar to that of polyvalent mannosylated ligands, which were
released from cell surfaces by the actions of very small molecules
(EGTA or EDTA) (37, 39).
The LBP that is bound to LPS aggregates may also contribute to their internalization by interacting with other cell surface molecules. Gegner et al. (5) showed that LBP is internalized with LPS as LPS-LBP-CD14 ternary complexes and proposed that weak interactions between LBP and other cell surface molecules stabilize these complexes. We recently reported that in the presence of LBP, LPS preferentially binds to mCD14 in low density, lipid-enriched plasma membrane domains (40). These domains may harbor the molecules that stabilize LPS-LBP-CD14 complexes and contribute to their rapid internalization.
Since native LPS is found in bacterial cell walls or membrane
fragments, where its physical state should be most similar to that of
highly aggregated LPS, the accelerating effect of aggregation on
internalization may be seen as a mechanism for cells to take up (clear)
(5) native LPS from their environment. Indeed, Grunwald and others (41)
have reported recently that CD14 can promote phagocytosis of
Gram-negative bacteria. The relationship between LPS aggregation state
and its ability to induce cellular responses is less clear. Others have
reported that the aggregation state of LPS can have a significant
influence on its stimulatory potency, with less aggregated forms (or
monomers) usually having greater activity (42, 43, 44, 45, 46). In these studies,
however, the cell stimulation assays were performed in the absence of
serum, so that the incubation mixtures lacked the transfer proteins
that promote interactions with mCD14. We found that aggregated,
partially disaggregated, and monomeric forms of LPS had approximately
equal stimulatory potencies under conditions that promote their binding
to mCD14 on THP-1 cells and monocytes (Table III). In keeping with
these observations, Hailman et al. (28) found that LPS-LBP aggregates
and monomeric LPS-sCD14 complexes made from R-form LPS (similar to the
LPS used in the present study) had similar potencies for stimulating
mCD14-dependent cytokine responses in cultured human macrophages and
neutrophils. Aggregated S-form LPS, however, was less stimulatory than
monomeric S-form LPS for reasons that are unclear. Experiments using
LBP mutants (47) or Abs (5) have dissociated signaling from bulk
binding of LPS aggregates, suggesting that the binding of aggregates to
mCD14 is not associated with cell stimulation. To reconcile these data,
one might conclude that mCD14 must monomerize LPS aggregates to some
extent to support signaling. LBP and sCD14 catalyze very rapid
monomerization of LPS, resulting in the formation of monomeric
LPS-sCD14 (28), and by inference, this process should also occur by the
actions of LBP and mCD14. This mechanism could explain why LBP is
required for sensitive responses to LPS aggregates (48, 49) but not for
LPS monomers (28).
It has been proposed that mCD14 may transfer LPS to the lipid bilayer
of the plasma membrane (50). We addressed this issue first by comparing
the time course of LPS internalization using the fluorescence quenching
and proteinase protection assays. As shown in Figure 3, these assays
gave virtually identical estimates of the relative proportions of
internal and surface LPS over time. We then measured surface-exposed
FITC-LPS before and after treating the cells with proteinase K or an
anti-fluorescein Ab. We found that protease treatment and Ab
quenching gave virtually identical estimates of surface FITC-LPS (Table II). Assuming that all surface-exposed FITC-LPS can be quenched by the
anti-fluorescein Ab, including FITC-LPS that is not susceptible to
removal by proteinase treatment (i.e., inserted into the lipid
bilayer), this result suggests that the LPS that remains on the cell
surface after incubation at 37°C is almost entirely protein bound. If
a large fraction of bound LPS is transferred to the lipid bilayer of
the cellular membrane(s) (50), this transfer probably occurs in an
internal compartment.
Although certain agonists can regulate their own internalization by
eukaryotic cells (51, 52, 53), stimulus-induced modulation of LPS
internalization has not been reported. LPS can down-regulate its own
catabolism by macrophages (21), however, and a recent report suggests
that LPS-induced signals may direct the intracellular traffic of
LPS-containing endocytic vesicles (18). Moreover, in keeping with
previous reports using other cell types (3, 4, 5), we found that the rate
at which LPS was internalized by THP-1 cells slowed significantly after
the first few minutes of exposure to LPS (Figs. 3 and 7). Does the
cellular response to LPS decrease its internalization rate? If so, one
would expect prior exposure to LPS to alter the rate at which newly
presented LPS is internalized. We found that pre-exposure to LPS for 5
min or 3 h or inhibition of LPS signals using a lipid A analogue
did not alter either the initial or the secondary rate of LPS
internalization (Fig. 8, A and B). In
addition, LPS-hyporesponsive and LPS-responsive murine macrophages
internalized LPS with virtually identical kinetics (Fig. 9). We
conclude, therefore, that LPS enters these cells by a constitutive
mechanism that is insensitive to its own stimulatory effects.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. R. L. Kitchens, Department of Internal Medicine, University of Texas Southwestern Medical Center, 5323 Harry Hines Blvd., Dallas, TX 75235–9113. E-mail:
3 Abbreviations used in this paper: mCD14, membrane-bound CD14; sCD14, soluble CD14; LBP, LPS-binding protein; PI-PLC, phosphatidylinositol-specific phospholipase C; RPMI-HB, RPMI 1640 with 10 mM HEPES (pH 7.4) and 0.3 mg/ml BSA; Ag-LPS, aggregated LPS; DAg-LPS, partially disaggregated LPS; HNEB, 20 mM HEPES (pH 7.4), 150 mM NaCl, 0.1 mM EDTA, and 0.3 mg/ml BSA; SFM, serum-free medium; NF-B, nuclear factor-B; MFI, mean fluorescence intensity; M-LPS, monomeric LPS.
Received for publication July 11, 1997. Accepted for publication October 29, 1997.
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,•) or empty vector (
and 
) were mixed with
increasing concentrations of [3H]LPS-sCD14 preformed
complexes (
) are shown. LPS bound in the absence of LBP is shown by open
symbols (
), internal LPS; 
) was prepared by
incubating LPS with sCD14 in the absence of LBP, centrifuging through a
100-kDa cut-off membrane, and diluting in RPMI 1640. DAg-LPS was
prepared by sonicating LPS in the presence of 0.1 mM EDTA (HNEB buffer)
and diluting in the same buffer (DAg-LPS, 
,
M-LPS; 
, DAg-LPS; •, Ag-LPS).