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(Journal of Leukocyte Biology. 2005;77:626-633.)
© 2005 by Society for Leukocyte Biology

Development and functional consequences of LPS tolerance in sinusoidal endothelial cells of the liver

Anja Uhrig*, Ramin Banafsche{dagger}, Michael Kremer{dagger}, Silke Hegenbarth{ddagger}, Alf Hamann§, Markus Neurath#, Guido Gerken, Andreas Limmer{ddagger} and Percy A. Knolle{ddagger},1

* Department of Transplant Surgery, University Mainz, Germany;
{dagger} Department of Surgery, University Heidelberg, Germany;
{ddagger} Institute of Molecular Medicine and Experimental Immunology, University Bonn, Germany;
§ Deutsches Rheumaforschungszentrum, Berlin, Germany;
Department of Gastroenterology and Hepatology, University Essen, Germany;
# 1st Medical Department, University Hospital, Mainz, Germany

1 Correspondence: Institute of Molecular Medicine and Experimental Immunology, University of Bonn, Sigmund-Freud-Str. 25, 53105 Bonn, Germany. E-mail: Percy.Knolle{at}ukb.uni-bonn.de


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Kupffer cells and liver sinusoidal endothelial cells (LSEC) clear portal venous blood from gut-derived bacterial degradation products such as lipopolysaccharide (LPS) without inducing a local inflammatory reaction. LPS tolerance was reported for Kupffer cells, but little is known whether sensitivity of LSEC toward LPS is dynamically regulated. Here, we demonstrate that LSEC react to LPS directly as a function of constitutive Toll-like receptor 4 (TLR4)/CD14 expression but gain a LPS-refractory state upon repetitive stimulation without loss of scavenger activity. LPS tolerance in LSEC is characterized by reduced nuclear localization of nuclear factor-{kappa}B upon LPS rechallenge. In contrast to monocytes, however, TLR4 surface expression of LSEC is not altered by LPS stimulation and thus does not account for LPS tolerance. Mechanistically, LPS tolerance in LSEC is linked to prostanoid production and may account for cross-tolerance of LPS-treated LSEC to interferon-{gamma} stimulation. Functionally, LPS tolerance in LSEC results in reduced leukocyte adhesion following LPS rechallenge as a consequence of decreased CD54 surface expression. Furthermore, LPS tolerance is operative in vivo, as we observed by intravital microscopy-reduced leukocyte adhesion to LSEC and improved sinusoidal microcirculation in the liver after repetitive LPS challenges. Our results support the notion that LPS tolerance in organ-resident scavenger LSEC contributes to local hepatic control of inflammation.

Key Words: liver • TLR-4 • lipopolysaccharide • cell adhesion


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
The liver is the central organ for removal of antigenic material from blood derived from the gastrointestinal tract (entering the liver with the portal vein) or from the systemic circulation (entering the liver via the hepatic artery) [1 , 2 ]. The anatomic position of the liver and the large amount of blood passing through human liver (≥2000 L per day) underline the importance of the liver as a filter system. Pathogenic, proinflammatory molecules such as lipopolysaccharide (LPS) enter the blood circulation from the gut and are cleared in the absence of any signs of inflammation by the hepatic sinusoidal cell populations of the reticulo-endothelial system [3 , 4 ], i.e., Kupffer cells and liver sinusoidal endothelial cells (LSEC), thus preventing their systemic distribution and development of widespread inflammatory reactions.

Hyporesponsiveness or tolerance to LPS has first been reported for monocytes and represents a host mechanism to limit tissue damage during systemic distribution of LPS [5 , 6 ]. A number of mechanisms are further reported to limit Kupffer cell reactivity to concentrations of LPS contained physiologically in portal blood. First, the unique hepatic microenvironment with its low arginin concentration lowers Kupffer cell sensitivity to LPS [7 ]. Second, LPS tolerance develops in monocytes through a blockade of intracellular signaling events and results in reduced expression of proinflammatory mediators upon LPS rechallenge, and Kupffer cells have been shown to develop LPS tolerance [8 ]. Third, Kupffer cells release interleukin (IL)-10 in response to LPS and thus control expression of proinflammatory mediators from other sinusoidal cell populations [9 ]. Along this line, IL-10 is involved in protection against LPS-induced liver damage [10 , 11 ].

The local mechanisms operative in precluding hepatic inflammation after LPS exposure with regard to hepatic endothelial cells have remained elusive. Here, we address the question of whether LSEC, which actively participate in removal of bacterial components such as LPS from the blood and are directly responsive to LPS stimulation, can autonomously control their response to LPS.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Isolation of murine LSEC
LSEC from mouse liver were isolated by centrifugal elutriation, as described previously [12 ]. Briefly, after portal-vein perfusion with Ca++-deprived buffer (0.05% collagenase A, Boehringer Mannheim, Germany), mechanically, dissection liver tissue was enzymatically digested to ensure further separation of cells. The cell suspension was passed through a sieve (300 µm) and washed twice with phosphate-buffered saline (PBS). Nonparenchymal cells were separated from parenchymal cells and debris by density-gradient centrifugation (1400 g) with metrizamide (1.089 g/cm3; Sigma Chemical Co., München, Germany). Nonparenchymal cells recovered from the gradient were further separated by counterflow centrifugal elutriation using an Avanti-JI centrifuge (Beckman, München, Germany) equipped with a JE-6B rotor and a standard elutriation chamber. The rotor speed was kept constant at 2500 rpm, and LSEC cells were recovered at a flow rate of 22 ml/min. LSEC were cultured at a density of 5 x 105 cells per well in collagen type I-coated tissue-culture dishes in Dulbecco’s modified Eagle’s medium/10% fetal calf serum. The purity of the LSEC fraction was tested by phagocytosis of opsonized sheep red blood cells (RBC; BAG-Hessen, Lich, Germany) and the typical appearance after incubation with DiI-acetylated low-density lipoprotein (AcLDL; Paesel and Lorei, Frankfurt, Germany), a substrate specific for endothelial cells. Purity of the isolated cell populations used for experiments was ≥98%. Following stimulation of LSEC cultures with LPS, we did not detect tumor necrosis factor {alpha} (TNF-{alpha}) or IL-10 in cell culture supernatants by specific sandwich enzyme-linked immunosorbent assay (ELISA). As Kupffer cells express TNF-{alpha} and IL-10 following LPS challenge [9 ], it is reasonable to assume that LSEC cultures, devoid of IL-10 or TNF-{alpha}, following LPS challenge, were not contaminated with Kupffer cells. Zymosan-Bodipy-FL (Molecular Probes, Leiden Netherlands) was incubated with LSEC or KC at a concentration of 10 µg/ml for 30 min.

Flow cytometry
Adherent LSEC were removed from solid support by incubation with Ca2+-free cell dissociation solution (Sigma Chemical Co.). LSEC (106) were stained with fluorescein isothiocyanate (FITC)-conjugated antibodies to CD54 or CD106 (3 µg/ml) at 4°C for 30 min. LSEC (104) were analyzed on a FACScan (Becton Dickinson, Heidelberg, Germany) using CellQuest software.

In vitro adhesion assay
LSEC were seeded into flat-bottom, 96-well plates (Falcon, Becton Dickinson) at a density of 105 cells/well. Splenic leukocytes were activated for 1 h with phorbol 12-myristate 13-acetate (15 ng/ml), washed extensively, and labeled with the fluorescent marker calcein-acetoxymethylester (Molecular Probes), according to the manufacturer’s instructions. Fluorochrome-labeled leukocytes (5x105) were incubated with mock-treated or LPS-stimulated LSEC (10 ng/ml for 18 h, Escherichia coli 055:B5, Sigma Chemical Co.). After 45 min at 37°C, unbound leukocytes were removed by gentle washing, and firmly adherent fluorescent leukocytes were detected by fluorescence microscopy. The relative change in numbers of leukocytes adherent to stimulated compared with mock-treated LSEC was calculated and is expressed in percent change of leukocyte adhesion to allow direct comparison of independent experiments.

Semiquantitative determination of gene expression in LSEC
Total RNA was extracted from LSEC using RNA Clean (AGS, Heidelberg, Germany) and subjected to reverse transcription (RT). Prior to first-strand cDNA synthesis, RNA was incubated for 10 min at 65°C. Total RNA was reverse-transcribed in a reaction mixture containing 5 µl transcription buffer (Gibco-BRL, Rockville, MD), 1 mmol/L dithiothreitol, 16 U RNase inhibitor (Promega, Madison, WI), 1 nmol random hexamer primers (Boehringer Mannheim), 4 mmol/L deoxy-unspecified nucleoside 5'-triphosphates (dNTPs; Boehringer Mannheim), and 200 U Moloney murine leukemia virus RT (Gibco-BRL) in a final volume of 25 µl. The reaction was performed at 40°C for 1 h, followed by a 10-min incubation at 65°C for enzyme inactivation. To allow semiquantitation, cDNA clones for murine CD54, murine CD106, and murine ß-actin were generated by polymerase chain reaction (PCR) using a TA cloning kit (Invitrogen, San Diego, CA). Primers were designed using Geneworks (Intelligenetics, Mountain View, CA) and sequence data from Genebank.

For semiquanitative PCR, A T7 promoter sequence (taatacgactcactatagggag) was added 5' to the 5' primers, to allow transcription of PCR products by T7 RNA polymerase. The following primers were used: ß-actin T7 5'GTGGGCCGCTCTAGGCACCA (428 bp product), 3'TAGCCCTCGTAGATGGGCACAG; CD54 5'TGCACGTGAACTGTTCTTCC (478 bp product), 3'CTGAGATCCAGTTCTGTGCG; CD106 5'AACTCCTTGCACTCTACTGCC (328 bp product), 3'GTTAGCTCCTTGCATTCAGC. The 5' primers were designed to bind to sequences within the respective cDNA clone. This ensured that hybridization conditions of the 5' primers containing the T7 promoter sequence were the same for LSEC cDNA and the respective cDNA clone.

The PCR reaction mixture was composed of 5 µl Taq buffer (Perkin Elmer, Norwalk, CT), 200 µmol/L dNTP, 200 mmol/L of each 3' and 5' primer, 2.5 U DNA polymerase (Perkin Elmer), and 5 µl cDNA solution in a total volume of 50 µl. Optimal annealing temperatures were determined to be 67°C for ß-actin and 61°C for CD54 and CD106. Thirty PCR cycles were run using the following conditions: cycle 1: 4'/94°C, 1'/X°C, 1'/72°C; cycle 2–29: 1'/94°C, 1'/X°C, 1'/72°C; cycle 30: 1'/94°C, 1'/X°C, 10'/72°C (X=annealing temperature).

As described previously, PCR products were transcribed into cRNA by T7 RNA polymerase [13 ]. Diluted PCR product (10 µl; 1:10) was added to 5 µl transcription buffer (200 mmol/L Tris-HCl, pH 8.0, 50 mmol/L MgCl2), 50 mmol/L ribonucleotide triphosphate solution containing adenosine 5'-triphosphate (ATP), cytidine-5'-triphosphate, and guanosine 5'-triphosphate (Boehringer Mannheim), 28 U RNase inhibitor (Promega), 34 U T7 RNA polymerase (Pharmacia, Uppsala, Sweden), and 6.3 µCi 35S-uridine 5'-triphosphate (Amersham, Braunschweig, Germany) in a total volume of 25 µl. The reaction mixture was incubated for 1 h at 37°C. Only one cRNA product per reaction was observed after separation by 6% polyacrylamide gel electrophoresis under denaturing conditions (8 mol/L urea, 40% formamide) and visualization by autoradiography (data not shown). Exact determination of incorporated radioactivity was achieved after acid precipitation and absorption of cRNA to glass fiber filters. Bound radioactivity was measured in a Beckman ß-counter. To correct for different RNA levels used in this assay and for variable efficiency in RT between different samples, the results obtained were normalized against ß-actin mRNA concentration.

Extraction of nuclear proteins and electrophoretic mobility shift assay (EMSA)
Nuclear proteins were isolated as described [14 ]. Oligonucleotides for EMSA were end-labeled with ({gamma}32P) ATP (>5000 Ci/mmol; Amersham, Arlington Heights, IL) using T4 polynucleotide kinase (New England Biolabs, Beverly, MA). The binding reaction was incubated at room temperature for 30 min. After electrophoresis, the gels were dried and exposed to Kodak MS films on intensifying screens at –80°C.

Intravital microscopy
Male Wistar rats, body mass 288 ± 12 g, were anesthesized with 20 mg/kg ketamine (Ketanest S®) intramuscularly and 25 mg/kg pentobarbital (Narcoren®) intraperitoneally and placed in a supine position; body temperature was maintained at 37.5 ± 0.5°C by means of a heating pad. The abdomen was opened by a subcostal transverse incision, and the left liver lobe was mobilized by minimal dissection of the falciform and triangular ligaments. The left liver lobe was partially exteriorized onto a suspension bench and covered with prewarmed Metocel® (isoosmotic Ringer-based gel). During intravital videomicroscopy, the animal remained on the heating pad, and continuous superfusion of warmed Ringer solution was on the exteriorized liver lobe.

Hepatic microcirculation was investigated with a modified Leica® video microscope (OPTO Inc., München, Germany). Leukocytes were stained with rhodamine 6G (Sigma Chemical Co.), and erythrocytes were labeled with FITC (Sigma Chemical Co.) to investigate leukocyte-endothelial interaction and sinusoidal perfusion, respectively. Filter combinations of 470 ± 20 nm/>515 nm (excitation/emission) and 545 ± 15 nm/>580 nm were used. Twenty-four nonadjacent hepatic acini and postsinusoidal venules were randomly selected per group and time-point and were recorded as described [15 ].

The following microcirculatory parameters were evaluated in off-line analysis, including sinusoidal perfusion rate (%), given as the percentage of nonperfused sinusoids of all observed sinusoids–a sinusoid was defined as perfused when flow of FITC-labeled RBC was detected; and numbers of permanently adherent leukocytes in the sinusoidal or postsinusoidal vascular segment, respectively–leukocytes were classified as permanently adherent when binding to the vessel wall exceeded 30 s. Data obtained were analyzed statistically as described [15 ].


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
LPS tolerance in LSEC
LSEC were isolated from mice by portal vein perfusion, mechanical dissociation, density centrifugation, and centrifugal elutriation. Purity of LSEC preparations was determined by flow cytometry using an endothelial cell- specific antibody (ME9F1) and uptake of the scavenger ligand AcLDL. Figure 1A demonstrates that more than 96% of cells used in experiments take up AcLDL and are positive for ME9F1. Moreover, confocal laser-scanning microscopy reveals mannose receptor-mediated uptake of ovalbumin into endosomal compartments in addition to typical scavenger receptor-mediated AcLDL uptake (Fig. 1B) . These results underline the most efficient scavenger function of LSEC and provide evidence that cells investigated here are LSEC and not macrophages. To further exclude the presence of contaminating Kupffer cells in LSEC preparations, we stained for the macrophage marker CD11b and performed phagocytosis assays. LSEC did not show expression of CD11b, whereas Kupffer cells were strongly positive (Fig. 1C) . Phagocytosis of Zymosan was only observed in Kupffer cells but not in LSEC, as demonstrated in Figure 1D .



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  		Figure 1. Characterization of LSEC, which were isolated from murine livers and taken into culture as described in Materials and Methods. (A) LSEC were incubated with 1 µg/ml Bodipy-FL-labeled AcLDL for 30 min and subsequently stained with ME9F1 antibody or an isotype control antibody. (B) LSEC were incubated with 1 µg/ml Bodipy-FL-labeled AcLDL and 5 µg/ml ovalbumin-Texas red for 30 min. Subcellular distribution of endocytosed ligand was determined by confocal laser-scanning microscopy. Original scale bar, 1 µm. (C) LSEC or KC were incubated with Zymosan-Bodipy-FL (10 µg/ml) for 30 min. Cells were subsequently washed, removed from the solid matrix, and isolated by flow cytometry. One out of two independent experiments is shown. Gray line, Untreated cells; black line, cells incubated with Zymosan. (D) LSEC or KC were stained with CD11b-phycoerythrin (PE) antibody and analyzed by flow cytometry. Gray line, LSEC; black line, KC.

 
Contact of LSEC with LPS in vitro leads to direct activation and release of the proinflammatory cytokine IL-6. LSEC responsiveness to LPS occurs in the absence of TNF-{alpha}, as neutralizing antibodies to TNF-{alpha} did not influence LPS-triggered IL-6 release from LSEC (Fig. 2A ). The phenotypic identification of LSEC in Figure 1 and the absence of LPS-induced TNF-{alpha} (data not shown) preclude that contaminating Kupffer cells in the primary cell populations were responsible for the observed reactivity toward LPS. Furthermore, LSEC sensitivity to LPS requires expression of functionally active TLR4, as LSEC from C3H-HeJ animals failed to release IL-6 in response to LPS stimulation (Fig. 2A) . Upon repeated LPS contact, responsiveness to LPS was diminished or lost (Fig. 2A) , suggesting that LSEC can autonomously control their reactivity to LPS. As LSEC constitutively express TLR4 (Fig. 2B) , we wondered whether LPS tolerance is associated with down-regulation of this receptor as described for monocytes/macrophages. However, surface expression of TLR4 in LSEC was not affected by contact with LPS, whereas J774 monocytes showed a clear reduction in TLR4 surface expression levels following exposure to LPS (Fig. 2B) . Similarly, TLR4 surface expression levels of Kupffer cells were not altered 4 h after contact with LPS (Fig. 2B) . Reduced nuclear translocation of transcriptionally active NF-{kappa}B is another prominent feature of LPS tolerance observed in monocytes/macrophages [5 ], which we observed in LPS-tolerant LSEC as well (Fig. 2C ; 3.5-fold difference by densitometric analysis). It is interesting that scavenger function of LSEC is not abrogated by previous contact with LPS. Uptake of fluorescently labeled LPS or a scavenger receptor ligand (AcLDL) was reduced but not prevented in LPS-treated versus mock-treated LSEC (Fig. 2D) .



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  		Figure 2. LPS tolerance in LSEC. (A) LSEC from C3H mice were stimulated with LPS (10 ng/ml) or LPS + neutralizing sheep antiserum to murine TNF-{alpha} or were pretreated with LPS (10 ng/ml) before LPS rechallenge 24 h later, and cell culture supernatants were assayed for IL-6 concentration by ELISA. LSEC from C3H-HeJ animals were used as control. Experiments were carried out in duplicates, and one out of three independent experiments is shown. TLR4, Toll-like receptor 4. (B) TLR4 surface expression levels of LSEC, J774 macrophages, and Kupffer cells were determined by staining with biotin-conjugated anti-TLR4 antibody (5 µg/ml) and detection with Streptavidin-PE. Shaded area, Isotype control; solid line, mock-treated cells; broken line, LPS-treated cells. (C) Nuclear translocation of nuclear factor (NF)-{kappa}B was determined by EMSA using equal amounts (15 µg) of nuclear proteins from endotoxin or mock-stimulated LSEC. NF-{kappa}B DNA-binding activity was assessed using a reference-binding site for NF-{kappa}B. The location of the specific retarded complex and the free probe is indicated. (D) Scavenger activity of LSEC was determined by uptake of substrates such as FITC-labeled LPS (10 µg/ml) or DiI-labeled AcLDL (50 µg/ml). Solid line, Mock-treated LSEC; faint line, LPS-treated LSEC; broken line, background fluorescence intensity of LSEC. Three independent experiments were performed. Down-regulation of scavenger receptor activity by LPS was not statistically significant by Student’s t-test.

 
Functional consequences of LPS tolerance in LSEC
We wondered whether LPS tolerance in LSEC contributed to the absence of hepatic inflammation during physiological encounter with gut-derived bacterial products. Therefore, we studied expression levels of adhesion molecules known to be important for interaction of LSEC with passenger leukocytes. Here, we report that LPS challenge leads to up-regulation of CD54 surface expression in naïve but not in LPS-tolerant LSEC (Fig. 3A ). Expression levels of other molecules relevant for leukocyte adhesion, i.e., CD106 as well as the chemokines monocyte chemoattractant protein-1 and IFN-inducible protein 10, were equally affected by LPS tolerance in LSEC (data not shown). To differentiate between reduced gene expression and increased internalization of surface molecules, we determined CD54 mRNA levels semiquantitatively by RT-PCR in naïve and LPS-tolerant LSEC upon LPS challenge. Figure 3B demonstrates that LPS-induced CD54 gene expression is reduced in LPS-tolerant LSEC, explaining the lack of increased surface expression. This observation is in line with reduced LPS-triggered nuclear translocation of NF-{kappa}B in LPS-tolerant LSEC (Fig. 2C) . Moreover, LPS-tolerant LSEC were cross-resistant to other proinflammatory stimuli and failed to up-regulate CD54 after IFN-{gamma} challenge (Fig. 3C) .



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  		Figure 3. LPS tolerance influences adhesion molecule expression in LSEC and leukocyte adhesion to LSEC. (A) LSEC were mock-treated or stimulated with LPS (10 ng/ml) in vitro for 18 h before rechallenge with LPS (10 ng/ml), and CD54 surface-expression levels were determined 18 h later by flow cytometry. Experiments were performed in duplicates, and 104 cells were analyzed per sample. Variation was below 10%. One representative out of three independent experiments is shown. (B) Relative CD54 gene expression levels (compared with ß-actin) following LPS challenge (10 ng/ml) were determined by semiquantitative RT-PCR of LSEC, LPS-pretreated (solid bars) or mock-treated (open bars). Results of three independent experiments are shown. (C) Cells were treated as described in A, but recombinant interferon-{gamma} (IFN-{gamma}; 25 IU/ml) was used instead of LPS for rechallenge. CD54 expression levels were determined by flow cytometry. One out of two independent experiments is shown. (D) Adhesion of leukocytes to LSEC in vitro was determined 18 h after LPS challenge (10 ng/ml) as described in Materials and Methods. Results from two experiments, carried out in triplicates, are shown with SD. (E) The time-course of leukocyte adhesion to LPS-challenged (10 ng/ml) LSEC is investigated. LPS-induced leukocyte adhesion to LPS-pretreated (solid bars) or mock-treated (open bars) LSEC was investigated. Results from two experiments, carried out in triplicates, are shown with SD.

 
In vivo studies using knockout mice have defined the central role of CD54 for hepatic leukocyte adhesion [16 ]. Using isolated LSEC, we determined in vitro the contribution of CD54 expression for leukocyte adhesion. Figure 3D shows that anti-CD54 antibodies reduced leukocyte adhesion to resting LSEC, supporting the notion that constitutive expression of CD54 is functional in LSEC-leukocyte interaction. Upon LPS challenge of LSEC in vitro, increased leukocyte adhesion was observed, which was blocked by anti-CD54 antibodies (Fig. 3D) . Accordingly, LPS challenge of LPS-tolerant LSEC, unable to up-regulate CD54 surface expression, failed to increase leukocyte adhesion (Fig. 3E) . These results suggest that LPS tolerance in LSEC is operative in attenuating CD54-dependent leukocyte adhesion during repetitive contact with LPS.

Effect of LPS tolerance on LSEC-leukocyte interaction in vivo
To investigate the role of LPS tolerance in leukocyte-LSEC interaction in vivo, we performed intravital microscopy to quantify leukocyte adhesion to hepatic sinusoidal endothelium. Rats were preconditioned with intravenous (i.v.) LPS or were mock-treated, and hepatic microcirculation was assessed upon re-exposure to i.v. LPS 24 h later by intravital microscopy. When rechallenged with LPS, preconditioned animals showed a significant reduction of permanently adherent leukocytes in hepatic sinusoids but not in hepatic venules (P≤0.01), suggesting distinct regulation of leukocyte adhesion in liver sinusoidal (Fig. 4A ) versus venular endothelium (Fig. 4B) . Moreover, the reduction of high-affinity leukocyte adhesion in the sinusoids was accompanied by improvement of the sinusoidal microcirculation, which was most pronounced 6 h after LPS challenge (11.1±4.8% in LPS-preconditioned animals vs. 34.8±12.1% nonperfused sinusoids/mm2 liver surface in mock-treated animals; Fig. 4C ). Although we cannot formally exclude a contribution of other hepatic cell populations to the results of in vivo experiments, it is clear that LPS tolerance is functional in controlling leukocyte adhesion to sinusoidal endothelial cells of the liver.



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  		Figure 4. Attenuation of LPS-induced leukocyte adhesion to hepatic sinusoidal endothelium and improvement of hepatic microcirculation after LPS preconditioning. Rats were i.v.-injected with LPS (100 µg/kg) or PBS prior to rechallenge with LPS (100 µg/kg) 24 h later. Intravital fluorescence microscopy was performed at 1, 2, 6, and 12 h after LPS rechallenge in anesthesized animals. Adhesion of leukocytes with high affinity was separately determined to sinusoidal endothelium (A) or postsinusoidal venular hepatic endothelium (B) by off-line video analysis. Representative pictures taken from videos at 1 h after LPS challenge are shown to the right of bar graphs. Leukocyte adhesion to postsinusoidal endothelium was quantified per mm2 endothelial surface, whereas adhesion to hepatic sinusoidal endothelium was quantified per mm2 liver surface. (C) The percentage of nonperfused hepatic sinusoids was determined in 24 nonadjacent hepatic acini and postsinusoidal venules per group and time-point.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
LSEC, as part of the reticulo-endothelial system, fulfill an important function as a scavenger cell population that clears portal venous blood from gut-derived pathogenic material such as LPS. Here, we demonstrate that repetitive LPS stimulation renders LSEC tolerant to further proinflammatory stimuli. LPS tolerance in LSEC develops independently of Kupffer cells and thus represents an endothelial cell autonomous control mechanism to prevent hepatic inflammation evoked by constant exposure to physiologic components of portal venous blood. It is important that scavenger function of LSEC is not abrogated by repetitive LPS contact (Fig. 2D) , which assures integrity of hepatic clearance function.

Moreover, LPS tolerance in LSEC has functional consequences beyond reduced release of proinflammatory mediators upon LPS rechallenge, as LPS-induced leukocyte adhesion in vitro is almost completely prevented in LPS-tolerant versus LPS-naïve LSEC (Fig. 3) . As leukocyte adhesion to LPS-stimulated and LPS-naïve LSEC in vitro was blocked by antibodies to CD54 (Fig. 3D) , and CD106 (data not shown) and LPS-tolerant LSEC fail to up-regulate CD54/CD106 expression in response to LPS challenge (Fig. 3B and 3C) , it is reasonable to assume that reduced, LPS-triggered leukocyte adhesion to LPS-tolerant LSEC is the result of low expression levels of adhesion molecules. Similar mechanisms are operative in vivo, where we observed reduced leukocyte adhesion specifically to sinusoidal endothelium but not to venular endothelium after repetitive LPS challenge (Fig. 4A and 4B) . Although we cannot formally exclude a role for LPS-sensitive Kupffer cells in this situation, it appears unlikely that Kupffer cells, located predominantly in the periportal area, released mediators that influenced leukocyte adhesion, as in that case, a periportal-perivenous gradient of leukocyte-adhesion should be present, but the contrary is observed. The clear, functional diversity between sinusoidal and venular endothelium in the liver rather argues for differences in intrinsic regulation of sensitivity to LPS in these two phenotypically and functionally different endothelial cell populations.

There are fundamental differences in the mechanisms mediating LPS tolerance in LSEC and monocytes, which rapidly down-regulate TLR4 surface expression upon LPS contact that renders them nonreceptive [17 ]; however, LSEC as well as Kupffer cells retain their constitutive TLR4 expression after LPS contact (Fig. 2B) . It is further possible that other adaptor molecules, such as myeloid differentiation protein-2, are differently expressed in LPS-pretreated LSEC and mediate resistance toward further stimuli [18 19 20 ]. LPS tolerance in LSEC rather seems to be mediated by endogenously produced prostanoids, as pharmacologic blockade of cyclo-oxygenase activity during initial LPS contact prevents development of LPS tolerance in LSEC (data not shown). Prostanoids produced by LSEC in response to LPS, such as prostaglandin E2 [21 ], may act via nuclear receptors to prevent gene expression induced by TLR4 ligand binding. Moreover, LPS-tolerant LSEC are cross-tolerant to other proinflammatory stimuli such as IFN-{gamma}, TNF-{alpha}, and even ischemia-reperfusion injury (data not shown). Thus, proinflammatory mediators released from Kupffer cells, liver-associated leukocytes, or passenger leukocytes are unlikely to have an effect on LPS-tolerant LSEC. LPS tolerance in LSEC may ensure an intact hepatic microcirculation and integrity of the hepatic endothelial cell layer by preventing binding and local activation of leukocytes during the physiologic situation.

However, LPS tolerance in LSEC is delicately balanced. LPS concentrations tenfold higher than during primary stimulation can overcome LPS tolerance in LSEC (data not shown), whereas LPS tolerance in macrophages is operative over a range of three to four orders of magnitude to prevent LPS-triggered cell stimulation [22 ]. The small dynamic range of LPS tolerance in LSEC may assure absence of inflammatory reactions during "physiologic" encounter but conversely, may allow reactivity of LSEC toward elevated LPS concentrations present during bacterial infection.

Although elimination of LPS from portal venous blood without induction of local hepatic inflammation is necessary for organ integrity, the presence of LPS appears to be relevant for the creation of the physiologic hepatic microenvironment. Komatsu et al. [23 ] reported that CD54 expression of LSEC is low-to-absent in germ-free mice but increases significantly upon bacterial colonization of the gastrointestinal tract. Our results demonstrate that LSEC, stimulated repetitively with LPS, show increased CD54 surface expression levels compared with naïve LSEC. This supports the notion that repetitive challenge with gut-derived LPS in vivo leads to a gradual increase of CD54 expression levels in LSEC. What are the functional implications of physiologic LPS exposure and constitutive expression of adhesion molecules on LSEC? We have previously reported that LSEC are similar to immature dendritic cells with respect to antigen-presenting cell function [24 ]. LSEC represent a unique, local organ-resident cell population capable of antigen presentation, which induces immune tolerance to exogenous antigens in CD4+ and CD8+ T cells. It is intriguing to speculate that constitutive adhesion molecule expression is operative in local hepatic induction of immune tolerance by enhancing interaction of passenger T cells with resident LSEC. Presentation of exogenous antigens on major histocompatibility complex type 1 (MHC I) molecules by LSEC is rather enhanced after contact of LSEC with LPS (A. Limmer et al., unpublished observation). Thus, LPS exposure increases in LSEC leukocyte adhesion and cross-presentation of exogenous antigens on MHC I molecules. It is interesting that evidence has been reported by a number of groups that immune tolerance against orally ingested antigens is linked to the presence of bacteria in the gut. Moreover, LSEC present orally ingested antigens efficiently, 2 h after feeding to T cells, and induce immune tolerance against oral antigens (A. Limmer et al., submitted). It is therefore conceivable that gut-derived LPS may contribute to hepatic induction of oral tolerance.


   ACKNOWLEDGEMENTS
 
This work was supported by grants from the Deutsche Forschungsgemeinschaft and the Volkswagenstiftung to P. A. K. A. U. and R. B. contributed equally.

Received June 9, 2004; revised November 18, 2004; accepted December 22, 2004.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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