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* Immunology Research Group, Department of Physiology and Biophysics, Faculty of Medicine, University of Calgary, Calgary, Canada; and
Division of Molecular Medicine, North Shore University/New York University School of Medicine, Manhasset, NY 11030
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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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200,000 people a year in
North America (1, 2). In these cases, it is the
inappropriate activation of inflammatory processes including
inappropriate leukocyte infiltration into tissues that causes the
progression to uncontrolled whole body inflammation (3).
It is thought that a major contributor to 1) localized infections as
well as 2) the morbidity associated with systemic infections is the
shedding of LPS from Gram-negative bacteria into the circulation
(4).
Leukocyte recruitment has generally been described as a multistep
cascade involving endothelial selectins (E- and P-selectin) and
leukocyte selectins (L-selectin) which permit the initial phase of
leukocyte recruitment, namely transient attachment to the endothelial
surface followed by leukocyte rolling along the vessel wall (5, 6). The second phase of leukocyte recruitment involves the
activation of integrins that mediate firm adhesion (5, 7).
In many local inflammatory conditions, including, e.g.,
ischemia/reperfusion of the mesentery or muscle, immunoneutralization
of one or more selectins completely inhibited leukocyte rolling and
therefore prevented leukocyte recruitment (8, 9, 10).
Interestingly, using identical antiselectin approaches to try to
inhibit LPS-induced leukocyte recruitment has proved to be far more
challenging. Although we were able to delay leukocyte rolling and
subsequent adhesion in mesentery with antiselectins, ultimately,
inhibition of one of the integrins (
4
integrin) was also necessary to prevent all leukocyte adhesion in
response to LPS (11). Further complexity was identified in
tissues like liver and lung where adhesion molecule inhibitor regimen
did not reduce leukocyte adhesion in response to LPS
(12, 13, 14).
Therefore, from a therapeutic viewpoint, due to the multitude of known
as well as to date unidentified adhesion mechanisms that are evoked
with LPS, antiadhesion therapy may not be a viable approach to limiting
leukocyte sequestration in endotoxemia. However, interruption of the
initiating signal perhaps at the level of the LPS receptor could
conceivably inhibit the multitude of adhesion molecules that are
activated and limit the untoward sequestration of leukocytes in the
lungs as well as other vascular beds. CD14 is considered the main LPS
receptor, present both as a soluble form in blood or as a
membrane-bound form in myeloid lineage cells. Indeed,
CD14-/- mice are at least 100 times more
resistant to LPS-induced mortality, and their macrophages also have
greatly reduced responsiveness as assessed by TNF- and IL-1
production (15). Because CD14 is a GPI-anchored protein
devoid of a transmembrane domain, by itself, it presumably cannot
transmit the activating signal into the cell. Indeed, in vitro work has
demonstrated that Toll-like receptor 4
(TLR4)3 activates the
immune system by functioning as a transmembrane coreceptor to CD14
(16, 17). Both TLR4-deficient
(TLR4d) mice and mice with a single point
mutation at aa 712 (proline to histidine) in the TLR4 gene
(C3H/HeJ) are resistant to the immunostimulatory and pathophysiological
effects of LPS (18, 19). Clearly, it seems reasonable to
hypothesize that altered leukocyte-endothelial cell interactions
observed in lungs and other tissues in response to LPS would be
entirely dependent on CD14 and associated TLR4.
Only a few studies have focused on the role CD14 and TLR4 in leukocyte sequestration. Based on initial work by Haziot et al. (15), CD14-deficient mice had reduced dissemination of Gram-negative bacteria. These investigators went on to discover that CD14-deficient mice as well as TLR4 mutant mice had an early and intense sequestration of neutrophils into the peritoneum, which was absent in wild-type mice. This was unexpected in light of the lack of responsiveness of these mutant mice to LPS. Therefore, we decided to systematically compare and contrast the role of CD14 and TLR4 in local and systemic LPS-induced leukocyte-endothelial cell interactions in vivo in numerous organs using two specialized techniques. First we used an in vivo assay system to visualize and study leukocyte behavior within a number of microvasculatures before and after LPS. Second, we used a sensitive, quantitative in vivo adhesion molecule expression system to examine endothelial responsiveness to LPS in all organs of CD14-/- and TLR4d mice. We show that TLR4d mice are completely resistant to LPS administration as it pertains to leukocyte-endothelial responses. By contrast, we identified that in some but not all microvascular beds partial endothelial responsiveness (protein expression and leukocyte function) to LPS was noted in CD14-/- mice.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Mice deficient in CD14 were generated by gene targeting in embryonic stem cells as previously described (15) and were backcrossed on a BALB/c background to the 10th generation. Wild-type BALB/c mice (CD14+/+) (The Jackson Laboratory, Bar Harbor, ME) were used as controls for the CD14-/- mice. TLR4d mice (C3H/HeJ mice) were obtained from The Jackson Laboratory. Wild-type C3H/HeN mice (TLR4+/+) (Charles River Laboratories, Montreal, Quebec, Canada) were used as controls for TLR4d mice. All mice weighed between 20 and 35 g and were between 6 and 10 wk of age at the time of use.
Intravital microscopy
Mice were anesthetized by i.p. injection of a mixture of xylazine hydrochloride (10 mg/kg; MTC Pharmaceuticals, Cambridge, Ontario, Canada) and ketamine hydrochloride (200 mg/kg; Rogar/STB, London, Ontario, Canada). The jugular vein was cannulated and used to administer additional anesthetic, fluorescent dyes, and various drugs. Animals were then prepared for viewing of either the skeletal muscle microcirculation (20) or the dermal microcirculation (21) as previously reported by our laboratory.
Cremaster muscle preparation. The cremaster muscle was dissected free of tissues and exteriorized onto an optically clear viewing pedestal. The muscle was cut longitudinally with a cautery and held flat against the pedestal by attaching silk sutures to the corners of the tissue. The muscle was then superfused with bicarbonate-buffered saline (pH 7.4).
Skin flap preparation. A midline dorsal incision was made. The skin was carefully separated from the underlying tissue, remaining attached laterally. Blood supply to the skin flap remained intact. The skin flap was then extended over a viewing pedestal, secured along the edges using 4-0 suture, exposing the dermal microvasculature. The exposed skin was continuously superfused with bicarbonate-buffered saline to avoid tissue dehydratation. Due to the thickness of the skin flap, leukocyte-endothelial cell interactions were not visible by transillumination. Therefore, for this protocol, animals were injected with the fluorescent dye, rhodamine 6G (0.3 mg/kg i.v.; Sigma-Aldrich, St. Louis, MO), immediately before microscopic visualization. Rhodamine 6G at the dose used labels leukocytes and platelets, allows detection of the same number of rolling leukocytes as transmitted light, and has no effect on leukocyte kinetics (22). Rhodamine 6G-associated fluorescence was visualized by epi-illumination at 510–560 nm using a 590-nm pore size emission filter (22).
The cremaster and dermal microcirculations were observed through an
intravital microscope with a x10 eyepice (Axioskop; Carl Zeiss Canada,
Don Mills, Ontario, Canada), a x25 objective lens for cremaster, and a
x40 objective lens for skin. A video camera (5100 HS; Panasonic,
Osaka, Japan) for the cremaster preparation and a fluorescent camera
(model C-2400-08; Hammamatsu Photonics, Hammamatsu City, Japan) for the
skin preparation were used to project the images onto a monitor, and
the images were recorded for playback analysis using a videocassette
recorder. Single unbranched venules (25–40 µm in diameter) were
selected, and to minimize variability the same section of the venule
was observed throughout the experiment. The number of rolling and
adherent leukocytes was determined off-line during video playback
analysis. Rolling leukocytes were defined as those cells moving at a
velocity less than that of erythrocytes within a given vessel.
Leukocyte rolling velocity was determined by measuring the time
required for a leukocyte to roll along a 100-µm length of venule.
Rolling velocity was determined for 20 leukocytes at each time
interval. Leukocytes were considered adherent to the venular
endothelium if they remained stationary for 30 s or longer.
Leukocyte emigration was defined as the number of extravascular
leukocytes per microscopic field of view and was determined by
averaging the data derived from four to five fields adjacent to
postcapillary venules. Venular diameter
(Dv) was measured on-line using a video
caliper (Microcirculation Research Institute, Texas A&M University,
College Station, TX). Centerline RBC velocity
(VRBC) was also measured on-line using
an optical Doppler velocimeter (Microcirculation Research Institute),
and mean RBC velocity (Vmean) was
determined as VRBC/1.6. Venular wall
shear rate (
) was calculated based on the Newtonian definition:
= 8
(Vmean/Dv)
(23).
Quantitation of endothelial activation
To determine the degree of endothelial activation two
endothelial adhesion molecules (P-selectin and VCAM-1) known to
contribute to leukocyte recruitment in endotoxemia were measured.
Expression of the adhesion molecules P-selectin and VCAM-1 were
quantified using a modified dual-radiolabeled Ab technique (21, 24). The Abs RB40.34 (against P-selectin) and 429 (MVCAM.A)
(against VCAM-1) were labeled with 125I. The Abs
A110-1 (a rat IgG1, 
isotype standard) and R35-95 (a rat IgG2a, 
isotype control Ig) were labeled with 131I. In
both cases, the Abs were labeled using the IodoGen method, as
previously described (21, 24). A110-1 and R35-95 were used
to detect nonspecific binding in the murine system.
To study P-selectin, animals were injected i.v. with a mixture of 10 µg 125I-anti-labeled P-selectin (RB40.34), and a variable dose of 131I-labeled nonbinding Ab (A110-1). To measure VCAM-1, mice were injected with 10 µg 125I-labeled anti-VCAM-1 (429 (MVCAM.A)), 25 µg unlabeled anti-VCAM-1 (429 (MVCAM.A)), and a variable dose of 131I-labeled nonbinding Ab (R35-95) calculated to achieve a total injected 131I activity of 400,000–600,000 cpm (total volume, 200 µl). This Ab combination was chosen after pilot experiments, conducted over a range of doses of unlabeled 429 (MVCAM.A), showed that this protocol ensured receptor saturation under stimulated conditions. In both cases, the Abs were allowed to circulate for 5 min; then the animals were heparinized. A blood sample was obtained from a carotid artery catheter; then the mice were exsanguinated by blood withdrawal through the carotid artery catheter and simultaneous i.v. infusion with bicarbonate-buffered saline. The lung, muscle, heart, brain, small bowel, large bowel, skin, pancreas, and liver were harvested and weighed. Both 131I and 125I activities were measured in plasma and tissue samples.
P-selectin and VCAM-1 expression was calculated per gram of tissue, by subtracting the accumulated activity of the nonbinding Ab (131I-labeled Ab) from the accumulated activity of the binding Ab (125I-labeled Ab). Data for P-selectin and VCAM-1 were presented as the percentage of the injected dose of Ab per gram of tissue. It has been previously demonstrated that this approach provides reliable quantitative values of adhesion molecule expression and that radiolabeled binding Ab can be displaced specifically with sufficient amounts of unlabeled Ab. The technique is sufficiently sensitive that very small, basal levels of P-selectin can be detected in wild-type mice relative to P-selectin-deficient mice where values are zero (24, 25).
Determination of tissue myeloperoxidase (MPO) activity
At the end of each experiment, samples of the lung were weighed, frozen on dry ice, and processed for determination of MPO activity. MPO is an enzyme found in cells of myeloid origin and has been used extensively as a biochemical marker of granulocyte (mainly neutrophil) infiltration into the lung (26, 27). The samples were stored at -20°C for no more than 1 wk before the MPO assay was performed. MPO activity was determined using an assay described previously, with the volumes of each reagent modified for use in 96-well microtiter plates (28). Change in OD450 during a 90-s period was determined using a kinetic microplate reader (Molecular Devices, Sunnyvale, CA).
Circulating leukocyte counts
At the end of each experiment, whole blood was drawn via cardiac puncture. Total leukocyte counts were performed, using a Bright-line hemocytometer (Hausser Scientific, Horsham, PA).
Experimental protocol
First, CD14-/-, TLR4d, and their respective wild-type mice (CD14+/+ and TLR4+/+) were prepared for intravital microscopy, and the microcirculations were examined for leukocyte-endothelial cell interactions under basal conditions. Circulating leukocyte counts as well as pulmonary MPO levels were also measured at the end of each experiment.
Local LPS administration.
In all experiments, LPS from Escherichia coli
0111:B4 dissolved in nonpyrogenic water was added to 0.2 ml saline.
This LPS is highly purified with <0.0008% of contaminating bacterial
proteins (15, 29). In these studies, LPS was administered
locally by s.c. injection beneath the scrotal skin using a 30-gauge
needle. Animals were returned to their cages for 3.5 h. The right
cremaster muscle was then prepared for intravital microscopy and
leukocyte-endothelial cell interactions, and hemodynamic parameters in
single postcapillary venules were examined. Preliminary experiments
indicated that local administration of 0.05 µg/kg LPS was optimal for
examination of leukocyte-endothelial interaction. To examine an
LPS-independent inflammatory response, the effect of local
administration of TNF- (0.5 µg) was examined in all mouse
strains.
Systemic (i.p.) LPS administration.
Mice received 0.5 mg/kg (12.5 µg/mouse) of the highly purified LPS
i.p. This dose was chosen to avoid any mortality over the study period
even after anesthesia. Moreover, preliminary dose-response studies
revealed that higher doses of LPS administered i.p. resulted in
stagnant peripheral microvasculatures. In one set of experiments, we
analyzed the expression of P-selectin adhesion molecule in different
mouse organs induced by LPS after 4 h treatment by using a
dual-radiolabeled Ab technique. To examine the peripheral
microvasculature directly, the dorsal skin or the cremaster muscle was
prepared for visualization by intravital microscopy and observed for 60
min between 3.5 and 4.5 h post-LPS administration. In all cases,
LPS-induced leukocyte sequestration was examined in
CD14-/-, TLR4d, and
their respective wild-type mice (CD14+/+ and
TLR4+/+).
Statistical analysis
All data are displayed as mean ± SEM. All data were analyzed using Student’s t test, and a Bonferroni correction was applied where multiple comparisons were necessary. A value of p < 0.05 was deemed significant.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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First we chose to examine a normal inflammatory response to local
LPS administration (0.05 µg/kg). This concentration of LPS had
absolutely no effect on hemodynamic parameters (data not shown) but
induced profound changes in leukocyte-endothelial cell interactions in
wild-type mice (Fig. 1
). In untreated
vessels of all four groups of mice (BALB/c (CD14+/+),
CD14-/-, C3H/HeN
(TLR4+/+), C3H/HeJ
(TLR4d)), 50–100 leukocytes can be seen
rolling through postcapillary venules every min (Fig. 1A).
These cells roll at relatively high velocities (40–80 µm/s) (Fig. 1B). Fewer than three cells are seen adhering (Fig. 1C) in the postcapillary venules. Local administration of
LPS into muscle significantly decreased leukocyte rolling velocity in
the muscle postcapillary venules of wild-type mice
(CD14+/+ and TLR4+/+),
suggesting activation of the local vasculature in these mice (Fig. 1B). Interestingly, absolutely no change in the rolling
velocity of leukocytes was noted in postcapillary venules of
CD14-/- and TLR4d mice
(Fig. 1B). The flux of rolling leukocytes was not altered by
the dose of LPS used in this study (Fig. 1A). Unlike local
LPS, one feature of systemic LPS is a profound drop in both the
circulating leukocytes and the flux of rolling leukocytes (later
section) in wild-type mice.
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also highlights that local administration of LPS led to an
increase of leukocyte adhesion (Fig. 1C) in wild-type mice
but not in CD14-/- and
TLR4d mice. No evidence of systemic effects was
noted in any strain of mice, because local LPS did not affect either
the number of circulating leukocytes or the number of neutrophils
sequestrated into the lungs (as assessed by MPO; data not shown).
Clearly, leukocyte adhesion was impaired in
CD14-/- and TLR4-/-
mice in response to local LPS administration.
To confirm that leukocyte-endothelial cell interactions can be induced
in CD14-/- and TLR4d
mice in response to other inflammatory mediators, TNF-, an
inflammatory mediator that acts independent of the LPS-CD14
interaction, was examined. Table I
reveals that intrascrotal injection of TNF- affected
leukocyte-endothelial cell interactions in
CD14-/-, TLR4d, and
their respective wild-type mice (CD14+/+ and
TLR4+/+) in a similar way. Local TNF- induced
in both mutant and wild-type mice a classical significant decrease
(80%) in the leukocyte rolling velocity and an increase in leukocyte
adhesion and emigration. Local administration of TNF- induced
similar changes in leukocyte rolling flux in
TLR4d and wild-type mice and in
CD14-/- and their respective wild-type mice
(data not shown).
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In preliminary data, we observed profound reductions
in blood flow using concentrations of LPS higher than 0.5 mg/kg i.p.
Therefore, we chose 0.5 mg/kg (12.5 µg/mouse) LPS, which induces
leukocyte sequestration without affecting hemodynamic parameters
including venular diameter, RBC velocity, and calculated wall shear
rates (Table II). Venules between 25 and
35 µm were chosen in all groups of animals. RBC velocity was
2.0–3.5 mm/s, and the calculated shear rate (based on velocity and
diameter) also did not increase or decrease with LPS administration
during the first 4 h. Therefore, hemodynamic changes were not a
confounding factor of leukocyte sequestration with systemic LPS
administration.
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We analyzed circulating leukocyte counts in
CD14+/+, CD14-/-,
TLR4+/+, and TLR4d mice
under baseline conditions. As shown in Fig. 2A, there are no differences
in baseline circulating leukocyte counts among the different mice.
Systemic LPS induced a profound drop in the number of circulating
leukocytes in wild-type mice (CD14+/+ and
TLR4+/+). Circulating lymphocytes, neutrophils,
and monocytes were decreased (data not shown). This decrease was
perhaps in part a result of a significant increase in the number of
leukocytes sequestered into the lungs as assessed by pulmonary MPO
activity (Fig. 2B). In contrast, neither
CD14-/- nor TLR4d mice
responded to systemic LPS, i.e., circulating leukocyte counts remained
unchanged in both groups of animals before and after LPS treatment
(Fig. 2A). The role of CD14 and TLR4 in leukocyte
sequestration in the lungs was examined because they are primary target
organs in sepsis. Lung MPO increased 2- to 3-fold in both strains of
animals in response to systemic LPS administration, whereas no response
was noted in CD14-/- and
TLR4d mice (Fig. 2B). The 8- to
10-U/mg tissue of MPO increase in lung suggests that the majority of
circulation neutrophils were sequestered in this organ.
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Leukocyte recruitment into peripheral tissues is
dependent on the ability of the endothelium to express selectins to
allow for leukocyte rolling followed by subsequent integrin-dependent
leukocyte adhesion. P-selectin expression was used as a marker of
endothelial activation in all vasculatures of wild-type and mutant
mice, because this molecule was previously shown to be an important
selectin in endotoxin-induced leukocyte sequestration (11, 14). Figs. 3 and 4 demonstrate a 20- to 50-fold
increase in P-selectin expression in all wild-type mouse tissues
(CD14+/+ and TLR4+/+)
tested. The organs are divided into CD14-dependent (Fig. 3) and
CD14-independent (Fig. 4) LPS responses. In
CD14-/- mice, there were no increases in
P-selectin expression in lung in response to LPS consistent with a lack
of leukocyte sequestration into this organ (Fig. 3A). No
increase in endothelial P-selectin expression was detected in response
to LPS in muscle, heart, brain, small bowel, or large bowel (Fig. 3, B–F). In surprising contrast, a consistent and
significant increase in P-selectin expression was observed in the skin
(4-fold) and pancreas (7-fold) of CD14-/- mice
(Fig. 4). Interestingly, all organs of TLR4d
mice remained completely unresponsive to LPS (Figs. 3 and 4). Although
the majority of leukocytes roll via P-selectin at 4 h LPS, a
second endothelial molecule (VCAM-1) was also elevated in
CD14-/- mice (Fig. 5). Again no increase in VCAM-1
expression in response to LPS was detected in
TLR4d mice, whereas an increase in VCAM-1
expression was detected in a number of tissues including pancreas (Fig. 5A), heart, liver, and large bowel (data not shown).
Surprisingly, a small amount of VCAM-1 was even detected in muscle and
lung, which could impact on mononuclear cell sequestration at later
times (Fig. 5, B and C) but not on the 4-h
leukocyte sequestration. Fig. 5D shows that some tissues,
e.g., small intestine, had no VCAM-1 up-regulation in
CD14-/- mice. However, VCAM-1 appeared to be
up-regulated in more tissues than P-selectin.
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To assess whether the organ-specific, differential adhesion
molecule responsiveness in different microvasculatures translated to
altered leukocyte function, we directly visualized leukocyte
trafficking in skin and muscle postcapillary venules using intravital
microscopy. LPS induced in both wild-type mice
(CD14+/+ and TLR4+/+), a
>90% reduction in leukocyte rolling flux within the skin (Fig. 6A), which may reflect our
work (Fig. 2) and previous reports of leukocytopenia (3, 13). By contrast, LPS did not alter the flux of rolling cells in
either CD14-/- or TLR4d
mice treated with LPS (Fig. 6A). The velocity of leukocyte
rolling is indicative of increased tether upon the leukocyte by
activated endothelium (expression of selectins and chemokines). A very
significant reduction in leukocyte rolling velocity was noted in
CD14+/+ mice (Fig. 6B). If LPS had no
impact in skin vasculature of CD14-/- mice, no
reduction in rolling velocity would be expected. However, in
CD14-/- mice, the velocity of rolling cells was
significantly reduced, indicating that the endothelium was activated in
CD14-/- skin postcapillary venules (Fig. 6B). No decrease in rolling velocity was seen in
TLR4d skin postcapillary venules. In the two
strains of wild-type mice, leukocyte adhesion increased 5- to 6-fold
(Fig. 6C). Most importantly, a significant increase in
leukocyte adhesion was noted in CD14-/- mice
(Fig. 6C) consistent with the increased adhesion molecule
expression in skin microvasculature. The latter responses were
completely absent in TLR4d mice (Fig. 6C) consistent with the view that TLR4 is absolutely
essential in LPS signaling even in tissues where CD14 was not
needed.
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Next, we examined leukocyte trafficking in the muscle
microvasculature wherein no endothelial activation (no P-selectin) was
detected in CD14-/- or
TLR4d mice stimulated with LPS. Under basal
conditions, 50–100 cells/min rolled through microvessels (Fig. 7A). By contrast after LPS
treatment, we noted that both CD14+/+ and
TLR4+/+ mice had a profound reduction in
leukocyte rolling flux within the cremaster (Fig. 7A),
similar to that seen in the skin. The velocity of leukocyte rolling,
which is indicative of local endothelial activation, was reduced by
95% in both wild-type mice strains after 4 h LPS administration
(Fig. 7B). In contrast, neither the flux of rolling cells
nor the velocity of these cells was altered in either
CD14-/- or TLR4d mice,
suggesting that the endothelium was not activated in the postcapillary
venules of these mice. It is interesting that in the muscle
postcapillary venules, very few cells ultimately adhered in response to
systemic LPS (Fig. 7C) in either wild-type or the mutant
mouse strains. Only a 2- to 3-fold increase in adhesion was noted
perhaps due to less effective synthesis of proinflammatory molecules
(chemokines, etc.) in the muscle vasculature. The same scale is used as
in Fig. 1C, highlighting the far more effective adhesion of
leukocytes in local (Fig. 1) than in systemic (Fig. 7) LPS. Regardless,
the data in Fig. 7 demonstrate significant changes in leukocyte
behavior (rolling, rolling velocity) in wild-type but not
CD14-/- mice in muscle, whereas responsiveness
was noted in skin (Fig. 6).
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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There are numerous reports to suggest that inhibition of adhesion molecules in lungs is not sufficient to prevent leukocyte accumulation into this tissue (12, 13). This is thought to be due to physical trapping of activated leukocytes within this microvascular bed due to the more narrow architecture of pulmonary capillaries (26, 27). Wang et al. (30) have reported that the endothelium increases in rigidity and perhaps contributes to the physical trapping of the general circulating pool of leukocytes. There may also be a direct increase in leukocyte rigidity in response to LPS and thereby increased sequestration within the pulmonary microvascular bed (30). Clearly, it is not surprising that some studies have reported only partial or no decrease in LPS-induced leukocyte sequestration with anti-adhesion therapy (13, 31). In this study, we for the first time have identified a genetic intervention (CD14-/- or TLR4d) that completely prevents any systemic LPS-induced leukocyte sequestration into the pulmonary microvasculature. Clearly, LPS stimulates the CD14 and TLR4 complex, which activates both leukocytes and endothelium to induce adhesion and/or physical trapping within the pulmonary microvasculature.
Although at least 10 TLRs have been identified, TLR2 and TLR4 have
received the most attention as recognition receptors of distinct
bacterial cell wall components and more specifically LPS (19, 32). Much evidence suggests that TLR4 is the dominant LPS TLR.
Initial work demonstrated that LPS stimulated NF-B-mediated gene
expression in HEK 293 cells only when those cells were transfected with
TLR4 cDNA (33). Mouse strains that lacked functional TLR4
(C3H/HeJ and C57BL/10ScCr) revealed resistance to LPS assessed as a
lack of cytokine production and no mortality (18). More
recently, TLR4-/- mice showed
hyporesponsiveness to LPS to an extent similar to that of C3H/HeJ mice
(19). Although TLR2 was initially also thought to function
as an LPS receptor, this was likely a result of contaminant found in
commercial LPS preparations (29, 34). Current evidence
argues against a major role for TLR2 in the physiological response to
LPS (19, 35, 36). For example,
TLR2-/- mice have normal cytokine profiles and
a similar degree of mortality in response to LPS (19). Our
study examined leukocyte-endothelial cell interactions directly with
highly purified LPS, and the data support the view that TLR2 is not
involved in LPS-induced leukocyte sequestration inasmuch as in the
absence of functional TLR4 there was no detectable sequestration of
leukocytes or endothelial activation. Clearly, our data are in
agreement that TLR4 is the only TLR involved in LPS-mediated
leukocyte-endothelial cell interactions.
Although the majority of work regarding TLR4 and CD14 has been performed in isolated macrophages, which are definitely activated during endotoxemia, it is quite likely that the early vascular responses to LPS occur via a direct effect of LPS on endothelium. Indeed, LPS will rapidly activate endothelium to induce synthesis of proadhesive molecules that sequester leukocytes independent of macrophages (37). Although endothelium also expresses TLR4 (38, 39, 40), unlike monocytes and neutrophils, endothelium is thought not to constitutively express membrane CD14 (41, 42, 43, 44). However, this cell will respond to LPS via a soluble form of CD14 (sCD14) present in blood (45, 46, 47, 48). Therefore, in the presence of plasma proteins, endothelium has all of the machinery necessary to rapidly respond to LPS and synthesize P-selectin, E-selectin, VCAM-1, and ICAM-1 as well as chemokines to induce leukocytes to roll, adhere, and ultimately emigrate into tissues. Our data would suggest that much like macrophages, endothelium responds to LPS exclusively via TLR4 to induce leukocyte sequestration.
Although this is to our knowledge the first report of LPS-induced leukocyte sequestration via a CD14-independent, TLR4-dependent pathway, a number of in vitro studies support this view. For example, Tsan et al. (49) reported that LPS was unable to induce TNF and manganese superoxide dismutase in isolated, CD14-/- peritoneal macrophages at low concentrations but could induce TNF and manganese superoxide dismutase at higher concentrations of LPS. Similar concentration-dependent findings have been reported in peritoneal macrophages for TNF message and for certain LPS-inducible genes including IL12 (p35 and p40) and COX-2 (50). Finally, in the original study by Haziot et al. (15), CD14-/- mice produced no cytokines at 20 mg/kg LPS but could produce IL-6 but not TNF at 200 mg/kg. To avoid concentration-dependent effects, we used 0.5 mg/kg LPS in this study (400-fold less), which did not induce cytokine production in CD14-/- mice (15) yet in the dermal microcirculation induced an increase in leukocyte-endothelium interactions. One could argue that, because we injected LPS i.p., CD14-/- peritoneal macrophages were exposed to relatively high concentrations of LPS, inducing release of cytokines that would have activated distal vasculatures. However, this seems very unlikely because 1) we used 400-fold less LPS than is required to induce cytokine production from CD14-/- macrophages, 2) no systemic signs of inflammation including a drop in circulating leukocytes were noted, and 3) the activation was site specific, i.e., occurred in skin but not muscle or lung.
Interestingly, TLR4d mice were resistant to LPS
responses that occurred independent of CD14. For example,
TLR4d mice were resistant to the increase in
P-selectin expression in all the tested organs; in the skin
microcirculation, TLR4d mice were completely
devoid of leukocyte effects in response to systemic LPS (Fig. 6).
Clearly, our data suggest a very sensitive second CD14-independent,
TLR4-dependent pathway of leukocyte sequestration. The selectivity of
this mechanism in skin rather than lung or muscle may reflect
LPS-binding proteins on dermal endothelium or extravascular cells
(dermal macrophages, fibroblasts, keratinocytes) that have evolved
alternative LPS detection systems.
Potential candidates as substitutes for CD14 include
2 integrins (CD11/CD18), the macrophage
scavenger receptor and L-selectin, which have all been proposed to bind
LPS and activate cells (51, 52, 53). However, to our
knowledge, none of these molecules is expressed in significant amounts
on endothelium. Very recently, heat shock proteins 70 and 90, chemokine
receptor 4, and growth differentiation factor 5 have been reported to
form a CD14-independent activation cluster after LPS ligation and are
involved in LPS signal transduction (54). However, no
mention of a need for TLR4 was made in that study. Finally, new reports
indicate that MD-2 is a genuine LPS-binding protein in a manner that is
independent of the assistance by either LPS-binding protein or CD14
(55, 56). Like CD14, MD-2 has no transmembrane domain and
apparently remains cell associated because it binds to TLR4
(57). Although extremely difficult to test in vivo, MD-2
may be a likely candidate to mediate significant CD14-independent,
TLR4-dependent leukocyte-endothelial cell responses to LPS. However,
this type of mechanism may be less likely to be organ specific, unless
different vasculatures display different levels of MD-2 and
TLR4.
In this study, we have for the first time described in vivo an LPS-induced leukocyte sequestration via a CD14-independent, TLR4-dependent pathway in skin. In addition, we have reported that the absence of CD14 or TLR4 prevents any leukocyte sequestration into the lungs and muscle. Thus, taken together, our results suggest that there are CD14-dependent and CD14-independent responses to LPS, but in both cases the responses are dependent on the presence of TLR4, suggesting that other proposed LPS receptors besides CD14 likely require TLR4 in the leukocyte sequestration response.
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2 Address correspondence and reprint requests to Dr. Paul Kubes, Department of Physiology and Biophysics, University of Calgary, 3330 Hospital Drive NW, Calgary, AB T2N 4N1 Canada. E-mail address: pkubes{at}ucalgary.ca
3 Abbreviations used in this paper: TLR4, Toll-like receptor 4; TLR4d, TLR4-deficient; MPO, myeloperoxidase.
Received for publication March 22, 2002. Accepted for publication June 21, 2002.
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H. Zhou, G. Andonegui, C. H. Y. Wong, and P. Kubes Role of Endothelial TLR4 for Neutrophil Recruitment into Central Nervous System Microvessels in Systemic Inflammation J. Immunol., October 15, 2009; 183(8): 5244 - 5250. [Abstract] [Full Text] [PDF]
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H. Zhou, B. M. Lapointe, S. R. Clark, L. Zbytnuik, and P. Kubes A Requirement for Microglial TLR4 in Leukocyte Recruitment into Brain in Response to Lipopolysaccharide J. Immunol., December 1, 2006; 177(11): 8103 - 8110. [Abstract] [Full Text] [PDF]
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S. C. Mullaly and P. Kubes The Role of TLR2 In Vivo following Challenge with Staphylococcus aureus and Prototypic Ligands J. Immunol., December 1, 2006; 177(11): 8154 - 8163. [Abstract] [Full Text] [PDF]
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, In
TLR4+/+, no rolling was seen at 4.5 h, but
small amounts of rolling were seen at early time points.