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Am J Physiol Gastrointest Liver Physiol 294: G520-G528, 2008. First published December 6, 2007; doi:10.1152/ajpgi.00489.2007
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LIVER AND BILIARY TRACT

Liver sinusoidal endothelial cells are the principal site for elimination of unfractionated heparin from the circulation

Cristina Ionica Øie,1 Randi Olsen,2 Bård Smedsrød,3 and John-Bjarne Hansen1

1Center for Atherothrombotic Research in Tromsø (CART), Department of Medicine, Institute of Clinical Medicine; 2Department of Electron Microscopy, Faculty of Medicine; 3Department of Cell Biology and Histology, Institute of Medical Biology, University of Tromsø, Tromsø, Norway

Submitted 25 October 2007 ; accepted in final form 4 December 2007


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
The mechanism of elimination of blood borne heparin was studied. To this end unfractionated heparin (UFH) was tagged with FITC, which served as both a visual marker and a site of labeling with 125I-iodine. UFH labeled in this manner did not alter the anticoagulant activity or binding specificity of the glycosaminoglycan. Labeled heparin administered intravenously to rats (0.1 IU/kg) had a circulatory t1/2 of 1.7 min, which was increased to 16 min upon coinjection with unlabeled UFH (100 IU/kg). At 15 min after injection, 71% of recovered radioactivity was found in liver. Liver cell separation revealed the following relative uptake capacity, expressed per cell: liver sinusoidal endothelial cell (LSEC)-parenchymal cell-Kupffer cell = 15:3.6:1. Fluorescence microscopy on liver sections showed accumulation of FITC-UFH only in cells lining the liver sinusoids. No fluorescence was detected in parenchymal cells or endothelial cells lining the central vein. Fluorescence microscopy of cultured LSECs following binding of FITC-UFH at 4°C and chasing at 37°C, showed accumulation of the probe in vesicles located at the periphery of the cells after 10 min, with transfer to large, evenly stained vesicles in the perinuclear region after 2 h. Immunogold electron microscopy of LSECs to probe the presence of FITC following injection of FITC-UFH along with BSA-gold to mark lysosomes demonstrated colocalization of the probe with the gold particles in the lysosomal compartment. Receptor-ligand competition experiments in primary cultures of LSECs indicated the presence of a specific heparin receptor, functionally distinct from the hyaluronan/scavenger receptor (Stabilin2). The results suggest a major role for LSECs in heparin elimination.

clearance; receptor-mediated endocytosis; fluorescence microscopy

HEPARIN IS A NATURALLY OCCURRING anticoagulant synthesized and secreted by connective tissue mast cells (15, 53). It is a mucopolysaccharide composed of alternating units of sulfated D-glucosamine and D-glucuronic acid. The esterified sulfuric acid component gives unfractionated heparin its acidic properties and electronegative charge. Unfractionated heparin (UFH) is heterogeneous with respect to molecular size, anticoagulant activity, and pharmacokinetic properties (30, 31). Even though UFH has been important for prevention and treatment of both venous and arterial thrombosis for decades, the mechanism by which UFH is eliminated from the circulation is largely unknown.

Studies in hepatectomized dogs (47) and patients with liver cirrhosis (67) showed increased half life of intravenous UFH in the circulation, indicating that the liver plays a crucial role in UFH removal. Both in humans (12) and in animal models (rats and rabbits) (5, 7), the plasma t1/2 of heparin increased with dose. Whole body autoradiography of rats injected with tritium-labeled heparin showed that the ligand was retained by the organs belonging to the reticuloendothelial system (RES), with the largest uptake in the liver (75). However, little is known about the cells involved in the clearance of UFH from the circulation. In 1939, Asplund et al. (1) observed heparin in the "sternzelln" of the liver sinusoids of rabbits, rats, and guinea pigs, but not in liver parenchymal cells (PCs) following injection of single and repeated doses of heparin. Furthermore, repeated administration of heparin into normal and atherosclerotic rabbits was succeeded by heavy accumulation of the polysaccharide in Kupffer cells (KCs) and in endothelial-like cells lining the liver sinusoid (28).

It is now well established that liver sinusoidal endothelial cells (LSECs) in mammals are the most important site for elimination of a variety of circulating physiological and nonphysiological soluble macromolecules (54, 64). LSECs exert their scavenger function by receptor-mediated endocytosis, thereby representing an important member of the hepatic RES together with the liver macrophages, the KCs (58). The purpose of the present study was to elucidate the role of LSECs in the clearance of UFH both in vivo and in vitro.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
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Experimental animals. Male albino rats, Sprague-Dawley (mean body wt 250 g), purchased from Scanbur (Sollentuna, Sweden), were kept under controlled animal room conditions at 21°C, relative humidity 55 ± 10% and 12:12 light-dark cycle (0800–2000 light) and fed a standard chow (Scanbur, Nittedal, Norway) ad libitum. For in vivo studies, anesthesia was induced with 4% Isofluran (Abbott Scandinavia, Solna, Sweden) and maintained at 2.1%. For in vitro studies, rats were anesthetized by subcutaneous injection of a mixture of Domitor, 0.4 mg/kg (Orion Pharma, Espoo, Finland) and Ketalar, 60 mg/kg (Pfizer, Lysaker, Norway). All experimental protocols were approved by the Norwegian Animal Research Authority in accordance with the Norwegian Animal Experimental and Scientific Purposes Act of 1986.

Chemicals. UFH (5,000 IU/ml) was obtained from Nycomed Pharma (Oslo, Norway). Carrier-free Na125I was from Perkin-Elmer Norge (Oslo, Norway), and 1,3,4,6-tetrachloro-3{alpha}, 6{alpha}-diphenylglycoluril (Iodogen) was from Pierce Chemical (Rockford, IL). Collagenase P was from Worthington Biochemical (Lakewood, NJ). Human serum albumin (HSA) was from Octapharma (Ziegelbrucke, Switzerland). Culture medium RPMI 1640, supplemented with 20 mM sodium bicarbonate, 0.006% (wt/vol) penicillin, and 0.01% (wt/vol) streptomycin, was from GIBCO BRL (Roskilde, Denmark). PBS was prepared as follows: 8.0 g/l NaCl, 0.2 g/l KCl, 2.9 g/l Na2HPO4·12 H2O, and 0.2 g/l KH2PO4. Fluorescein isothiocyanate (FITC), bovine serum albumin (BSA), mannan (MANN), chondroitin 4-sulfate (CS-A), and chondroitin 6-sulfate (CS-C) were from Sigma (St. Louis, MO). Sephadex G-25 (PD-10 columns) and Percoll were from Amersham Biotech (Uppsala, Sweden). Acetylated LDL (Ac-LDL) was from Biomedical Technologies, Stoughton, MA. Formaldehyde-treated bovine serum albumin (FSA) was prepared as described (45). Purified human IgG and high-molecular-weight hyaluronan (HA) were from Pharmacia, Uppsala, Sweden. Anti-Stabilin2 antibody (anti-rS2) was prepared as described (44). Single collagen {alpha}-chains (COLLA) were obtained by incubation of native triple helical collagen (Vitrogen, Palo Alto, CA) for 60 min at 60°C. Human IgG (10 mg/ml) was heated for 30 min at 63°C to prepare aggregated IgG (AGG) (32). Advanced glycation end products-modified bovine serum albumin (AGE-BSA) was prepared as described (29).

Immunoreagents. FITC-UFH was localized on thawed cryosections by use of specific monoclonal FITC antibodies (18.6 g/l) from Boehringer Mannheim Biochemica, Mannheim, Germany. Rabbit anti-mouse IgG from Cappel Research Products (Durham, NC) was used as a bridging antibody. Protein A-gold was purchased from Cell Microscopy Center, Department of Cell Biology, University Medical Center Utrecht, The Netherlands. Colloidal gold (5 nm diameter) was prepared by the tannic acid method and coupled to 25 mg/ml BSA as described (55, 56).

Labeling procedures. UFH (25 mg/ml) and BSA (0.5 g) were labeled with FITC as described (26). To remove unbound FITC, the solutions were run on a PD-10 column, equilibrated, and eluted with phosphate-buffered saline (PBS). When compared with the unlabeled, starting material, the anticoagulant activity (as measured by the STA-Rotachrom Heparin kit from Diagnostica Stago, Asnieres-sur-Seine, France) of FITC-UFH was unchanged. FITC-UFH and FSA (0.1 mg/ml) in 0.1 ml PBS were labeled with carrier-free Na125I in a direct reaction employing Iodogen as oxidizing agent, as described (42). Radiolabeled FITC-UFH and FSA and free iodine were separated by gel filtration on a PD-10 column equilibrated with PBS containing 1% HSA. The labeling method gave a specific radioactivity of 3–5 x 106 cpm/µg for both FITC-UFH and FSA. Owing to the photosensitivity of halogenated fluorescein, great care was taken to protect FITC-labeled UFH from light.

Blood clearance and in vivo anatomical distribution. Blood clearance and anatomical distribution of intravenously administered 125I-FITC-UFH were determined as described (59). Briefly, trace amounts of 125I-FITC-UFH (~0.1 IU/kg) alone, or together with unlabeled UFH in concentrations of 1, 20, and 100 IU/kg in 0.5 ml physiological saline, were injected into a lateral tail vein. Immediately thereafter, blood samples of 25 µl were collected from the tail tip into calibrated capillary tubes. The rate of elimination from the circulation was determined by relating the radioactivity measured in all blood samples to the first sample (1 min), which was set to 100%. Anatomical distribution of 125I-FITC-UFH was assessed 15 min after intravenous administration of 1 IU/kg 125I-FITC-UFH alone or together with 100 IU/kg of unlabeled UFH. The abdomen was cut open and the liver washed free of blood with 200 ml of physiological saline. The following organs were analyzed for radioactivity: liver, spleen, kidneys, stomach, intestines, urine, heart, eyes, brain, muscle, and blood. The total radioactivity in blood was calculated by using the fact that rats contain 5.75 ml blood per 100 g body wt (13).

Hepatocellular distribution. At 15 min after intravenous administration of 125I-FITC-UFH (1 IU/kg), a cannula was inserted into the portal vein and collagenase perfusion and purification of liver cells carried out as described below. The amount of radioactivity per million cells was measured in suspensions of PCs, LSECs, and KCs solubilized in 1% SDS. Cell numbers were assessed by visual counting in a phase-contrast microscope. The uptake per cell was calculated on the basis of the fact that the ratio between the number of KCs, LSECs, and PCs in rat liver is 1:2.5:7.7 (50). The method for determining the hepatocellular distribution of different ligands was previously used by us and by others (46, 59, 61, 70, 71).

Isolation of PCs, KCs, and LSECs from rat liver. The method for preparation of pure cultures of functionally intact PCs, KCs, and LSECs from a single rat liver has been described elsewhere (62). In short, the liver was perfused with collagenase, and the resulting single cell suspension was subjected to velocity and density centrifugations in Percoll gradients to produce purified suspensions of PCs and nonparenchymal cells (NPCs). The NPC suspension was a mixture of KCs and LSECs, and essentially devoid of PCs, red blood cells, and debris. The NPC suspension was seeded in 25-cm2 culture dishes (Nunk, Roskilde, Denmark) coated with BSA treated with 1% glutaraldehyde. Following a 30-min incubation at 37°C, only KCs attached and spread onto the substrate. Unattached cells, mainly LSECs, were transferred to 2-cm2 culture dishes coated with 0.1% collagen to enable attachment and spreading of these cells. All in vitro experiments were performed on freshly isolated cells, within the first 24 h. Serum is not required for primary cultures of LSECs, as the cells can be maintained only in RPMI 1640 for at least 24 h (63). Moreover, serum was omitted from the culture medium to avoid heparin degradation by serum heparitinase (49) and interactions between heparin and serum proteins.

Fluorescence microscopy of liver. At 15 min following intravenous administration of 200 IU of FITC-UFH in 0.5 ml of physiological saline, the liver was washed free of blood. Small cubes (~0.5 cm3) were randomly cut out of the liver tissue and snap frozen in isopentane in liquid nitrogen. Cryostat sections were fixed with acetone and mounted on coverslips. The specimens were examined with an Axiophot photomicroscope (Carl Zeiss, Oberkochen, Germany). Photomicrographs were taken with a Nikon DS 5MC digital camera.

Intracellular transport. Primary cultures of rat LSECs, established on 14-mm-diameter glass coverslips coated with 0.1% collagen, were incubated overnight at 4°C on a rocking platform with 0.2 ml of RPMI 1640 containing 0.1 mg/ml FITC-UFH. After removal of the unbound ligand by washing three times with PBS, bound ligand was chased for 0, 1, 5, 10, 20, 30, 60, and 120 min in fresh, prewarmed culture medium at 37°C. Incubations were terminated by fixation with 4% paraformaldehyde, 0.5% glutaraldehyde in PBS, and embedding in antifade medium consisting of 1 g DABCO (Sigma) in a solution of 5 ml PBS and 5 ml glycerol. The specimens were examined as described above.

Preparation of specimens for electron microscopy examination. Rats were injected with BSA-gold (5-nm diameter) in 0.5 ml physiological saline 10 min before the injection of 200 IU FITC-UFH in 0.5 ml physiological saline. Liver perfusion and isolation of LSECs was performed 30 min after the last injection. Cultures of LSECs (5 x 106), seeded in 12 cm2 dishes coated with 0.1% collagen, were incubated for 4 h at 37°C with RPMI 1640 containing 0.1 mg/ml FITC-UFH. Intracellular transport was terminated by washing the cultures with ice-cold PBS to eliminate excess of FITC-UFH, followed by fixation with 4% formaldehyde and 0.5% glutaraldehyde in PBS overnight. After quenching in 0.12% glycine, the cells were gently detached from culture dishes and pelleted in 12% gelatin by a 30-s centrifugation at 12,000 g. The cell pellet was placed in 2.3 M sucrose overnight, mounted on cryo pins, and frozen by immersion in liquid nitrogen.

Immunocytochemistry. Ultrathin sections of frozen pellets of LSECs were obtained by using a Leica EM UC6 ultramicrotome with a Leica FC6 cryochamber (Leica Mikrosysteme, Vienna, Austria) and a Diatome immuno-diamond knife (Diatome, Bile, Switzerland) at –120°C. The sections were retrieved in a mixture (50:50) of 2.3 mol/l sucrose and methylcellulose (34) and transferred to carbon-coated grids. Immunocytochemical labeling was performed as previously described (17) by using a working dilution of 1:1,500 anti FITC antibody-fish skin gelatin. Antibodies were detected by protein A-gold complexes. Double labeling was performed in a sequential manner, using a fixative block (1% glutaraldehyde in water) as described (57). The sections were dried and examined in a JEM 1010 transmission electron microscope (JEOL) operating at 80 kV, and micrographs were obtained with a Morada 11 Mega pixel camera.

Endocytosis experiments. Primary cultures of LSECs established in 2-cm2 dishes coated with 0.1% collagen were incubated for 2 h at 37°C with 0.2 ml RPMI 1640 containing trace amounts of 125I-FITC-UFH (~0.005 IU; 25 ng) alone (control), or together with excess amounts of unlabeled UFH, FITC-UFH, FSA, HA, AGE-BSA, CS-A, CS-C, Ac-LDL, FITC-BSA, COLLA, MANN, and AGG (0.1 mg/ml). For the experiments using the antibody to Stabilin2, cultures of LSECs were incubated for 30 min at 37°C with 0.2 ml RPMI 1640 containing 0.1 mg/ml anti-rS2, followed by 2-h incubation with trace amounts of 125I-FITC-UFH. Experiments were terminated by collecting the supernatant along with one washing volume of 0.5 ml PBS. Cell-associated 125I-FITC-UFH (sum of surface-bound ligand and endocytosed ligand) was quantified by measuring the amount of label solubilized in 1% SDS. Serum has been shown to be a rich source of proteins and other molecules that can inhibit the uptake via the scavenger receptors in LSECs (20). Therefore serum was omitted from the medium. On the basis of the knowledge that endocytosis represents the most important physiological function of LSECs, we used endocytosis of 125I-FSA each time an in vitro experiment was performed to test the integrity and the functionality of the cells (14).

Statistical analysis. The statistical analyses were performed with the SPSS statistical package for Windows version 14.0 (SPSS, Chicago, Ill). The GLM for analysis of variance was used to assess dosage effects. Differences in hepatocellular distribution were performed by an unpaired Student's t-test. Statistical analysis of the half-life data was performed using Graph Pad Prism 4. Differences were considered to be statistically significant if two-sided P values were less than 0.05. Results represent three separate experiments if not otherwise stated. Results are expressed as means ± SD.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Blood clearance and anatomical and hepatocellular distribution. Elimination of heparin from the circulation was investigated by injecting trace amounts of 125I-FITC-UFH (0.1 IU/kg) through the tail vein of rats (Fig. 1). The kinetics of clearance followed a monoexponential decay curve with a half-life of 1.71 min. To determine whether the 125I-FITC-UFH clearance was specific, increasing doses of unlabeled UFH were administered along with the radiolabeled tracer. The results showed that unlabeled UFH competed with 125I-FITC-UFH for clearance in a dose-dependent manner. Doses of 1, 20, and 100 IU/kg of unlabeled UFH resulted in plasma half-lives of 125I-FITC-UFH of 3.23, 7.37, and 16.28 min, respectively (P < 0.001). Approximately 18% of the recovered radioactivity was still present in the blood circulation 15 min after administration of trace amounts of 125I-FITC-UFH and stayed at a constant level for at least 1 h (not shown).


Figure 1
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  		Fig. 1. Removal of 125I-FITC-unfractionated heparin (UFH) from the circulation. 125I-FITC-UFH (0.1 IU/kg in 0.5 ml physiological saline) was administered intravenously either alone ({circ}) or along with increasing doses of unlabeled UFH [1 IU/kg ({triangleup}), 20 IU/kg (bullet), and 100 IU/kg ({blacktriangleup})] through the tail vein, and radioactivity was measured in blood samples over time. The percentage of ligand remaining in circulation was calculated, taking radioactivity in the blood sample drawn at 1 min after injection as 100%. Each data point represents mean ± SE (n = 4 for 0.1 IU/kg and n = 3 for 1, 20, and 100 IU/kg UFH). The solid lines represent the best fit of the experimental data describing 1-phase exponential clearance (R2 = 0.96 for 0.1 IU/kg, R2 = 0.98 for 1 IU/kg, R2 = 0.92 for 20 IU/kg, and R2 = 0.97 for 100 IU/kg).

 
To investigate the anatomical distribution of 125I-FITC-UFH, various organs were surgically removed and radioactivity was measured 15 min after intravenous administration of radiolabeled ligand into a lateral tail vein. Approximately 71% of the recovered radioactivity was found in the liver, 16% was found in the blood, and the rest (13%) was evenly distributed as minor amounts in other tissues and organs (Fig. 2).


Figure 2
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  		Fig. 2. Anatomical distribution of 125I-FITC-UFH. 125I-FITC-UFH (0.1 IU/kg in 0.5 ml physiological saline) was administered intravenously through the tail vein of rats, and radioactivity in organs was measured after 15 min. The results are presented as percentage of total recovered radioactivity. Radioactivity in tissues and organs other than those shown in the figure was less than 1%. Bars are means ± SD for 3 separate experiments. L, liver; B, blood; Int, intestines; K, kidneys; S, spleen.

 
Fluorescence microscopy. Fluorescence microscopy of random liver sections taken from tissue cubes fixed 15 min after intravenous administration of FITC-UFH revealed that the fluorescence accumulated only in cells lining the liver sinusoids, whereas no fluorescence was found in PCs or central vein endothelial cells (Fig. 3).


Figure 3
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  		Fig. 3. Phase-contrast (A) and fluorescence micrograph (B) of rat liver cryosections. At 15 min following intravenous administration of 200 IU of FITC-UFH in 0.5 ml physiological saline, the liver was washed free of blood. Small cubes (~0.5 cm3) were randomly cut out of the liver tissue and snap frozen in isopentane in liquid nitrogen. Cryostat sections were fixed with acetone, mounted on coverslips, and examined by fluorescence microscopy. The fluorescence accumulated exclusively in cells lining the liver sinusoids (arrowheads), and not in parenchymal cells (P) or central vein (CV) endothelial cells (arrows). Scale bar, 50 µm.

 
The observation that FITC-UFH was taken up by the liver prompted us to investigate the distribution of radiolabeled FITC-UFH in the different cell populations of the liver. The liver cells were isolated 15 min after intravenous administration of 125I-FITC-UFH (1 IU/kg), and the radioactivity per million cells in the different cell populations was measured. On the basis of relative distribution of cells in the rat liver (50), it could be calculated that the relative uptake capacity per cell was 15:3.6:1 for LSEC, PC, and KC, respectively. A similar hepatocellular distribution was found when an excess amount of unlabeled UFH (100 IU/kg) was injected along with the radiolabeled FITC-UFH (Fig. 4).


Figure 4
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  		Fig. 4. Hepatocellular distribution of heparin. At 15 min after intravenous administration of 125I-FITC-UFH (1 IU/kg, open bars) alone or together with 100 IU/kg (shaded bars) of unlabeled UFH, the liver cells were dispersed by collagenase perfusion, and the parenchymal cells (PCs), sinusoidal endothelial cells (LSECs), and Kupffer cells (KCs) were isolated. The amount of radioactivity per million cells was measured in suspensions of PCs, LSECs, and KCs solubilized in 1% SDS. Cell numbers were assessed by visual counting in a phase-contrast microscope. The uptake per cell was calculated based on the knowledge that the ratio between the number of KCs, LSECs, and PCs in rat liver is 1:2.5:7.7 (50). Bars are means ± SD for 3 separate experiments.

 
Intracellular transport. To study the intracellular transport of heparin, 0.1 mg/ml of FITC-UFH was incubated with primary cultures of LSECs overnight at 4°C to allow binding to receptors expressed at the cell surface. The cultures were then washed to remove unbound ligand, and transferred to 37°C to allow internalization and intracellular transport. Figure 5 shows the dynamics of intracellular transport of endocytosed FITC-UFH at 37°C. After binding at 4°C overnight, the FITC-UFH was diffusely located over the entire cell surface. After 10 min of subsequent chase at 37°C, the diffuse surface staining was changed to a pointlike pattern, located in the periphery of the cells and in the perinuclear area. The punctuate staining grew in size to form ring structures that appeared brighter after 30 min of chase. After 60 min the ring structures changed to become smaller in size and more perinuclearly located. The concentration of the probe continued to render these perinuclear small vesicles even brighter after 120 min of chase.


Figure 5
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  		Fig. 5. Morphological pulse-chase study on intracellular transport of endocytosed FITC-UFH. Cultures of LSECs were pulsed overnight with FITC-UFH (0.1 mg/ml) at 4°C. Chasing was performed after removal of unbound ligand by washing with PBS and transferring the cultures to 37°C. The cultures were fixed with 4% formaldehyde-0.5% glutaraldehyde in PBS after chase periods of 0, 10, 30, 60, and 120 min as indicated and examined by fluorescence microscopy. At chase start the probe was diffusely distributed over the cell surface. After 10 min the probe was concentrated in vesicles of varying size and location. The endocytic vesicles containing the probe grew in size until 30 min, at which time they reached their maximal size. Note that the probe appears only along the peripheral aspects of the larger vesicles (arrows), giving them a ringlike appearance. After 60 min and up to 120 min, the probe is concentrated in smaller, evenly stained perinuclear vesicles. Scale bar, 10 µm.

 
Ultrastructural studies. To determine the ultrastructural aspects of the late phase of intracellular transport of endocytosed FITC-UFH in LSECs, a known marker for the lysosomal compartment, BSA-gold (27, 52), was administered intravenously in rats 10 min before the administration of FITC-UFH. At 30 min after injection of the probes LSECs were isolated and prepared for immunoelectron microscopy. The presence of FITC was probed with anti-FITC-protein A gold (10 nm) complexes. Colocalization of FITC (10 nm gold) with the lysosomal marker BSA-gold (5 nm) strongly suggested that FITC-UFH was present in lysosomes (Fig. 6). As a negative control, BSA-gold was injected alone into rats. This procedure gave exactly the same localization of this lysosomal marker as when it was administered along with FITC-UFH (results not shown).


Figure 6
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  		Fig. 6. Electron micrograph of LSEC cultures. BSA-gold, a lysosomal marker, was administered intravenously 10 min prior to administration of 200 IU FITC-UFH. Isolation of LSECs was performed 30 min after the injection of FITC-UFH. The isolated cells were subsequently incubated for 2 h at 37°C before fixation with 4% paraformaldehyde-0.5% glutaraldehyde in PBS. The micrograph shows an LSEC cryosection labeled with anti-FITC visualized with protein A-gold (10 nm) that colocalized with BSA-gold (5 nm) in the lysosomes. Scale bar, 1 µm.

 
Specificity of endocytosis. The specificity of endocytosis of FITC-UFH in primary cultures of LSECs was studied by attempting to inhibit the uptake of 125I-FITC-UFH, using excess amounts (0.1 mg/ml) of FITC-UFH, native, unlabeled UFH, and other ligands known to be taken up by the scavenger receptors in LSECs (Fig. 7). The presence of FITC-UFH and native UFH inhibited the uptake of 125I-FITC-UFH by 94 and 93%, respectively. So far, three specific receptors for endocytosis have been identified and characterized on LSECs, i.e., the HA/scavenger receptor (Stabilin2), the mannose/collagen {alpha}-chain receptor (MANN/COLLA-R), and the Fc-{gamma} receptor (36, 38, 40, 44). Because heparin has the highest negative charge density of any known biological macromolecules (9), we speculated that heparin would be recognized by the scavenger receptor, which is a group of different endocytosis receptors with only one feature in common, namely the ability to take up negatively charged macromolecules (58). To settle this question, competition studies for endocytosis of 125I-FITC-UFH were carried out, using FSA, HA, AGE-BSA, Ac-LDL, CS-A, and CS-C (6, 16, 33, 43, 60, 66). The results (Fig. 7) show that these ligands only slightly inhibited the uptake of 125I-FITC-UFH, with HA as the most potent inhibitor (23%). Moreover, the antibody to Stabilin2, the receptor protein responsible for the endocytic clearance of HA and CS from lymph fluid and blood (25, 78), did not inhibit the uptake of 125I-FITC-UFH by LSECs. The observation that FITC-BSA did not inhibit the uptake of 125I-FITC-UFH suggests that the FITC adduct proper did not mediate the binding of FITC-UHF to LSECs. As negative controls we used MANN, COLLA, and AGG, ligands known to be taken up by the MANN/COLLA-R, and the Fc-{gamma} receptor in LSECs. No inhibition was observed by any of these three ligands.


Figure 7
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  		Fig. 7. Specificity of endocytosis. Cultures of 1 x 106 LSECs were incubated for 2 h at 37°C with trace amounts of 125I-FITC-UFH (~25 ng) alone (Control) or together with excess amounts of unlabeled UFH or FITC-UFH (0.1 mg/ml), as well as with excess amounts of formaldehyde treated bovine serum albumin (FSA), hyaluronan (HA), advanced glycation end products-modified bovine serum albumin (AGE-BSA), chondroitin 4-sulfate (CS-A), chondroitin 6-sulfate (CS-C), acetylated LDL (Ac-LDL), anti-rS2, FITC-BSA, collagen {alpha}-chains (COLLA), mannan (MANN), or aggregated IgG (AGG) (0.1 mg/ml). Cells were incubated with anti-rS2 30 min at 37°C prior addition of 125I-FITC-UFH. Cell-associated radioactivity was quantified by measuring the amount of label solubilized in 1% SDS. Uptake in control cultures was set to 100%. The results of each experiment, presented as a percent of control value, are means of 3 experiments run in triplicate. Bars are means ± SD.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
Our findings provide evidence for a major role of LSECs in the elimination of heparin. Even though the liver has been recognized as a major organ responsible for elimination of heparin, the precise mechanism by which heparin is eliminated from the circulation is largely unknown. Most studies on distribution of heparin in liver performed in vitro using fractionated heparin report that the KCs and the PCs are the main cells for uptake (19, 48, 74, 76), whereas only two studies indicated uptake in LSECs as well (28, 51). LSECs are an important site for elimination of a variety of circulating physiological and nonphysiological soluble macromolecules. The cells perform their scavenger function by receptor-mediated endocytosis (54, 64). Against this background, we decided to study the elimination of blood-borne heparin using both in vivo and in vitro experimental models and to determine the role of LSECs in the elimination of UFH. We found a dose-dependent increase in serum half-life; ~70% of recovered 125I-FITC-UFH was found in the liver. Fluorescence microscopy of liver sections following in vivo administration of FITC-UFH showed accumulation of the probe exclusively in cells lining the liver sinusoids. Liver cell separation showed that the radioactivity was mainly found in LSECs. Pulse-chase studies in primary cultures of LSECs showed accumulation of the probe in large, densely stained vesicles in the perinuclear region. With time, the probe colocalized with a marker of the lysosomal compartment, as assessed by electron microscopy. Endocytosis studies in cultured LSECs showed that the uptake of 125I-FITC-UFH was inhibited by surplus amounts of unlabeled UFH, whereas other ligands for the scavenger receptors inhibited the uptake only modestly.

Most previous studies have used iodinated (3, 65) or tritium-labeled heparin (11, 73, 77) for metabolic studies of heparin and assessment of organ distribution by autoradiography and recovered radioactivity in isolated organs. However, FITC-labeled UFH has also been shown to be suitable for studying heparin distribution and metabolism (11). In our study, we decided to use FITC-labeled UFH given a number of advantages with this complex. FITC-UFH, which is prepared by a simple one-step "mix-and-incubate" step, is easy to label with 125I, resulting in a stable radiolabeled complex with high specific radioactivity, allowing metabolic studies and organ distribution to be determined by simple measurement of gamma-radiation in isolated tissues and cells. Since FITC is trapped intralysosomally after internalization of FITC-conjugated ligand in the target cells, it can be traced by fluorescence microscopy, and it may also be detected by gold-labeled anti-FITC antibodies at the electron microscopic level (27). An objection to FITC-labeling is the potential effect on the physiochemical properties of heparin, and thereby its pharmacokinetic properties. However, several observations in our study provide evidence against such an objection. First, the anticoagulant activities of UFH and FITC-UFH, as measured by its anti-Xa activity, were the same. Second, circulatory elimination of trace amounts of 125I-FITC-UFH was competitively inhibited by increasing concentrations of unlabeled UFH, as shown by an increased serum half-life. Third, endocytosis of 125I-FITC-UFH in primary cultures of LSECs was almost completely inhibited by unlabeled UFH and FITC-UFH, providing further evidence for a specific and identical site for binding and internalization of native UFH and FITC-UFH. Fourth, the dose-dependent elimination of 125I-FITC-UFH in our study was similar to that reported by 125I-UFH in a rabbit model (7).

Previous studies have shown that intravenously administered heparin attached to vascular endothelium (39). This binding was assumed to contribute significantly to the rapid phase of the UFH clearance. This assumption was further supported by experimental studies on vascular endothelial cells in vitro, showing a slow binding to saturable and specific binding sites at the endothelial surface. Moreover, the capacity for internalization of heparin was reported to be very low in these cells (2, 4, 72). In our study, fluorescence microscopy of liver sections fixed 15 min after administration of FITC-UFH in vivo showed no fluorescence in vascular endothelium lining the central vein, whereas strong staining was observed along the cells lining the liver sinusoids. Our findings thus indicate that the vascular endothelium in the liver appears not to contribute much to the binding of UFH.

In agreement with previous studies (12, 35), we found that intravenously administered UFH accumulated mainly in the liver. Studies in primary cultures of KCs and PCs from rats reported a specific binding and uptake of fractionated heparin of low molecular weight (7,000 Da) and high molecular weight (20,000 Da) (74, 76). To our knowledge, the uptake of unfractionated heparin by LSECs was mentioned in two studies (28, 51). Only one of these (51) reported a relative LSEC-PC-KC uptake capacity per cell of 3.6:2.4:1 after injection of ~300 IU/kg 35S-labeled heparin. In our study, the relative LSEC-PC-KC uptake capacity per cell was 15:3.6:1 after injection of 125I-FITC-UFH. The differences between the studies may be due to the different methods used for liver perfusion and purification of the LSECs. The protocol used by us for LSECs isolation and purification is mild and rapid (with most steps at 4°C) and interferes minimally with the integrity of the cells. Moreover, the marker adduct FITC, carried into the cell after endocytosis of FITC-UFH, is trapped in lysosomes of the target cells. As 125I is attached to FITC molecules, the radioactivity taken up in the cells remains trapped inside the cells during isolation, purification, and cultivation of the cells. In the study mentioned above (51), 30 min after the injection of 35S-heparin, the LSECs were isolated by means of selective enzymatic destruction of PCs, by mincing the liver in pronase for 60 min at 37°C. The use of pronase could trigger rupture of LSECs cellular membrane and thus release of the radioactivity. The time period used between killing the animal and preparing the LSECs (2 h) and the physiological temperature (37°C) used during LSEC preparation, in addition to the 30 min in vivo exposure of the animal to the injected heparin, could also lead to desulfation and/or degradation of the 35S-heparin, leading to the release of the 35S-label from LSECs. This could thus give the impression of reduced heparin uptake by LSECs. In our study, the relative distribution of heparin in liver cells was similar with administration of low (1 IU/kg) or high (100 IU/kg) heparin doses, suggesting that LSECs have a high capacity for removing UFH from the circulation.

To preserve as many as possible of the in vivo features of the cells, it is essential that they can be used for experiments soon after the start of the isolation procedure, and that they can be kept in culture for longer periods without losing their viability. Therefore we used primary cultures of LSECs for all in vitro studies.

The following observations suggest that heparin is removed from the circulation by receptor-mediated uptake in LSECs. First, removal of 125I-FITC-UFH from the circulation was inhibited by excess amounts of unlabeled UFH. This finding indicates an actual competition for a specific binding site with limited capacity. Second, studies in primary cultures of LSECs showed that the uptake of 125I-FITC-UFH was inhibited efficiently by excessive amounts of unlabeled UFH and FITC-UFH, whereas only a minor inhibition was observed by other ligands for receptor-mediated endocytosis. HA was found to have the highest inhibitory effect (23%). Previously, similar inhibition of 125I-HA uptake by heparin was reported in cultured rat LSECs (25), and the authors suggested that the receptor for endocytosis of HA present in LSECs did not recognize heparin. The inability of heparin to inhibit the uptake of HA (and vice versa) has also been reported in vivo (18). The fact that the HA receptor [HARE; also called Stabilin2 and FEEL-2 (24)], does not recognize UFH was confirmed by our study when we showed that the antibody to Stabilin2 had no inhibitory effect on the endocytosis of 125I-FITC-UFH by LSECs. However, the receptor for HA may only partially recognize UFH, since HA inhibited the uptake of UFH by only 23%. Third, morphological pulse chase studies in vitro using FITC-labeled UFH revealed accumulation of the probe in intracellular vesicles, suggesting that the ligand was internalized by primary cultures of LSECs. These studies showed subsequent transport of FITC-UFH to two distinct intracellular compartments, with circular doughnut shapes appearing after 10 min, changing to perinuclear vesicles over the next 2 h. Similar structures were previously described during internalization of NH2-terminal propeptide of type I procollagen (PINP) and denatured collagen in rat LSECs (26, 46), ligands shown to be removed from the blood circulation by LSECs by receptor-mediated endocytosis. Fourth, simultaneous intravenous administration of FITC-UFH and BSA-gold particles followed by LSECs isolation and examination of ultrathin cryosections labeled with anti-FITC antibodies showed colocalization of the probe with the BSA-gold particles in the lysosomes. These observations suggest that internalized UFH travels through the endocytic compartments of LSECs, with final accumulation in lysosomes.

It is well known that UFH has the ability to bind to a number of plasma proteins due to the interactions between the negatively charged sulfate groups of UFH and basic amino acid residues of the proteins (10). We believe that this could explain why 18% of the recovered 125I-FITC-UFH was still present in the blood 1 h after injection. Therefore it is reasonable to assume that heparin-protein complexes formed in the circulation may affect the protein elimination. Heparin is known to exert its anticoagulant activity by forming heparin-antithrombin complexes that accelerate antithrombin-mediated inhibition of its target proteases (i.e., factor Xa and thrombin) (8), and, by mobilizing tissue factor pathway inhibitor (TFPI) from vascular endothelium into the circulation, forming heparin-TFPI complexes (69). Prolonged infusion of intravenous UFH is known to promote a pronounced decrease in circulating antithrombin (21, 41) and TFPI (22) accompanied by an increase in thrombotic events immediately after discontinuation of heparin infusion (68). The decrease in plasma antithrombin during continuous UFH treatment has been proposed to occur due to increased elimination of heparin-antithrombin complexes exceeding the rate of synthesis in the liver (21, 67). The latter assumption is supported by studies showing that patients with liver cirrhosis did not have the expected decrease in plasma antithrombin during heparin elimination (67). Furthermore, later studies reported increased synthesis and release of TFPI in vascular endothelial cells by UFH (23, 37), supporting the concept that increased elimination of heparin-TFPI complexes in the liver is the underlying mechanism for depletion of intravascular TFPI during heparin treatment.

In conclusion, our experimental studies in vivo and in vitro provide strong evidence that LSECs are the principal site for binding and uptake of UFH. Receptor-ligand competition studies, along with the use of specific anti-receptor antibodies, showed that the receptor for endocytosis of heparin in LSECs is distinct from the HA/scavenger receptor (Stabilin2). Formation of heparin-protein complexes during heparin treatment may have important clinical implications by affecting the elimination of the proteins. Further studies are warranted to elucidate the specific mechanism for binding and uptake of heparin in these cells, as well as identification and characterization of the heparin binding protein responsible for the heparin recognition and internalization in LSECs.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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REFERENCES
 
This study was supported by an independent grant from Pfizer AS and by a grant from The Norwegian Research Council (B. Smedsrød).


   ACKNOWLEDGMENTS
 
The authors wish to thank Merethe A. Lorentzen, Department of Pathology, University Hospital of Northern Norway, for excellent preparation of liver sections; Rod Wolstenholme, audiovisual department, for preparing and sizing the figures; and Dr. Peter A. G. McCourt for proofreading the manuscript.

C. I. Øie designed and performed the research, collected, analyzed, and interpreted data; performed statistical analysis; and wrote the manuscript. R. Olsen performed the immunohistochemistry and electron microscopy studies. B. Smedsrød and J-B. Hansen were responsible for funding, supervised the overall project, and contributed to writing the manuscript. All authors checked the final version of the manuscript.


   FOOTNOTES
 

Address for reprint requests and other correspondence: C. I. Øie, Center for Atherothrombotic Research in Tromsø, Dept. of Medicine, Institute of Clinical Medicine, Univ. of Tromsø, N-9037 Tromsø, Norway (e-mail: cristina{at}fagmed.uit.no)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 

  1. Asplund J, Borell U, Holmgren H. Untersuchungen über die Speicherung des Heparins im Tierorganismus sowie über seine Resorptionsmöglichkeiten in Darm und Placenta. Z Mikrosk Anat Forsch 46: 16–67, 1939.
  2. Barzu T, Molho P, Tobelem G, Petitou M, Caen J. Binding and endocytosis of heparin by human endothelial cells in culture. Biochim Biophys Acta 845: 196–203, 1985.[Medline]
  3. Barzu T, Molho P, Tobelem G, Petitou M, Caen JP. Binding of heparin and low molecular weight heparin fragments to human vascular endothelial cells in culture. Nouv Rev Fr Hematol 26: 243–247, 1984.[Medline]
  4. Barzu T, van Rijn JL, Petitou M, Tobelem G, Caen JP. Heparin degradation in the endothelial cells. Thromb Res 47: 601–609, 1987.[CrossRef][Medline]
  5. Bjornsson TD, Levy G. Pharmacokinetics of heparin. II. Studies of time dependence in rats. J Pharmacol Exp Ther 210: 243–246, 1979.[Abstract/Free Full Text]
  6. Blomhoff R, Drevon CA, Eskild W, Helgerud P, Norum KR, Berg T. Clearance of acetyl low density lipoprotein by rat liver endothelial cells. Implications for hepatic cholesterol metabolism. J Biol Chem 259: 8898–8903, 1984.[Abstract/Free Full Text]
  7. Boneu B, Caranobe C, Gabaig AM, Dupouy D, Sie P, Buchanan MR, Hirsh J. Evidence for a saturable mechanism of disappearance of standard heparin in rabbits. Thromb Res 46: 835–844, 1987.[CrossRef][Web of Science][Medline]
  8. Bourin MC, Lindahl U. Glycosaminoglycans and the regulation of blood coagulation. Biochem J 289: 313–330, 1993.[Web of Science][Medline]
  9. Capila I, Linhardt RJ. Heparin-protein interactions. Angew Chem Int Ed Engl 41: 391–412, 2002.
  10. Cardin AD, Weintraub HJ. Molecular modeling of protein-glycosaminoglycan interactions. Arteriosclerosis 9: 21–32, 1989.[Abstract/Free Full Text]
  11. Castellot JJ Jr, Wong K, Herman B, Hoover RL, Albertini DF, Wright TC, Caleb BL, Karnovsky MJ. Binding and internalization of heparin by vascular smooth muscle cells. J Cell Physiol 124: 13–20, 1985.[CrossRef][Web of Science][Medline]
  12. Dawes J, Papper DS. Catabolism of low-dose heparin in man. Thromb Res 14: 845–860, 1979.[CrossRef][Web of Science][Medline]
  13. Dittmer DS. (editor; analysis and compilation by Altman PL). Blood and Other Body Fluids. Washington, DC: FASEB, 1961.
  14. Elvevold K, Nedredal GI, Revhaug A, Bertheussen K, Smedsrod B. Long-term preservation of high endocytic activity in primary cultures of pig liver sinusoidal endothelial cells. Eur J Cell Biol 84: 749–764, 2005.[CrossRef][Web of Science][Medline]
  15. Enerback L, Kolset SO, Kusche M, Hjerpe A, Lindahl U. Glycosaminoglycans in rat mucosal mast cells. Biochem J 227: 661–668, 1985.[Medline]
  16. Eskild W, Kindberg GM, Smedsrod B, Blomhoff R, Norum KR, Berg T. Intracellular transport of formaldehyde-treated serum albumin in liver endothelial cells after uptake via scavenger receptors. Biochem J 258: 511–520, 1989.[Medline]
  17. Griffiths G, Burke B, Lucocq J. Fine Structure Immunocytochemistry. Berlin: Springer-Verlag, 1993, p. 265–270.
  18. Gustafson S, Bjorkman T. Circulating hyaluronan, chondroitin sulphate and dextran sulphate bind to a liver receptor that does not recognize heparin. Glycoconj J 14: 561–568, 1997.[CrossRef][Medline]
  19. Haba M, Urano K, Yuasa H, Watanabe J. Molecular weight dependency in the uptake of fractionated [3H]heparin in isolated rat Kupffer cells. Biol Pharm Bull 19: 864–868, 1996.[Web of Science][Medline]
  20. Hansen B, Melkko J, Smedsrod B. Serum is a rich source of ligands for the scavenger receptor of hepatic sinusoidal endothelial cells. Mol Cell Biochem 229: 63–72, 2002.[CrossRef][Medline]
  21. Hansen JB, Sandset PM. Differential effects of low molecular weight heparin and unfractionated heparin on circulating levels of antithrombin and tissue factor pathway inhibitor (TFPI): a possible mechanism for difference in therapeutic efficacy. Thromb Res 91: 177–181, 1998.[CrossRef][Web of Science][Medline]
  22. Hansen JB, Sandset PM, Huseby KR, Huseby NE, Nordoy A. Depletion of intravascular pools of tissue factor pathway inhibitor (TFPI) during repeated or continuous intravenous infusion of heparin in man. Thromb Haemost 76: 703–709, 1996.[Web of Science][Medline]
  23. Hansen JB, Svensson B, Olsen R, Ezban M, Osterud B, Paulssen RH. Heparin induces synthesis and secretion of tissue factor pathway inhibitor from endothelial cells in vitro. Thromb Haemost 83: 937–943, 2000.[Web of Science][Medline]
  24. Harris EN, Kyosseva SV, Weigel JA, Weigel PH. Expression, processing, and glycosaminoglycan binding activity of the recombinant human 315-kDa hyaluronic acid receptor for endocytosis (HARE). J Biol Chem 282: 2785–2797, 2007.[Abstract/Free Full Text]
  25. Harris EN, Weigel JA, Weigel PH. Endocytic function, glycosaminoglycan specificity, and antibody sensitivity of the recombinant human 190-kDa hyaluronan receptor for endocytosis (HARE). J Biol Chem 279: 36201–36209, 2004.[Abstract/Free Full Text]
  26. Hellevik T, Bondevik A, Smedsrod B. Intracellular fate of endocytosed collagen in rat liver endothelial cells. Exp Cell Res 223: 39–49, 1996.[CrossRef][Medline]
  27. Hellevik T, Martinez I, Olsen R, Toh BH, Webster P, Smedsrod B. Transport of residual endocytosed products into terminal lysosomes occurs slowly in rat liver endothelial cells. Hepatology 28: 1378–1389, 1998.[CrossRef][Medline]
  28. Hiebert L. The uptake of heparin by liver sinusoidal cells in normal and atherosclerotic rabbits. Thromb Res 21: 383–390, 1981.[CrossRef][Medline]
  29. Ito Y, Sorensen KK, Bethea NW, Svistounov D, McCuskey MK, Smedsrod BH, McCuskey RS. Age-related changes in the hepatic microcirculation in mice. Exp Gerontol 42: 789–797, 2007.[CrossRef][Medline]
  30. Johnson EA, Mulloy B. The molecular-weight range of mucosal-heparin preparations. Carbohydr Res 51: 119–127, 1976.[CrossRef][Web of Science][Medline]
  31. Kandrotas RJ. Heparin pharmacokinetics and pharmacodynamics. Clin Pharmacokinet 22: 359–374, 1992.[Web of Science][Medline]
  32. Kijlstra A, Van Es LA, Daha MR. Enhanced degradation of soluble immunoglobulin aggregates by macrophages in the presence of complement. Immunology 37: 673–680, 1979.[Medline]
  33. Laurent TC, Fraser JR, Pertoft H, Smedsrod B. Binding of hyaluronate and chondroitin sulphate to liver endothelial cells. Biochem J 234: 653–658, 1986.[Web of Science][Medline]
  34. Liou W, Geuze HJ, Slot JW. Improving structural integrity of cryosections for immunogold labeling. Histochem Cell Biol 106: 41–58, 1996.[CrossRef][Web of Science][Medline]
  35. Losito R, Barlow G, Lemieux E. 3H-heparin and antithrombin III in the isolated liver perfusion. Thromb Res 10: 83–93, 1977.[CrossRef][Medline]
  36. Lovdal T, Andersen E, Brech A, Berg T. Fc receptor mediated endocytosis of small soluble immunoglobulin G immune complexes in Kupffer and endothelial cells from rat liver. J Cell Sci 113: 3255–3266, 2000.[Abstract]
  37. Lupu C, Poulsen E, Roquefeuil S, Westmuckett AD, Kakkar VV, Lupu F. Cellular effects of heparin on the production and release of tissue factor pathway inhibitor in human endothelial cells in culture. Arterioscler Thromb Vasc Biol 19: 2251–2262, 1999.[Abstract/Free Full Text]
  38. Magnusson S, Berg T. Extremely rapid endocytosis mediated by the mannose receptor of sinusoidal endothelial rat liver cells. Biochem J 257: 651–656, 1989.[Web of Science][Medline]
  39. Mahadoo J, Heibert L, Jaques LB. Vascular sequestration of heparin. Thromb Res 12: 79–90, 1978.[CrossRef][Web of Science][Medline]
  40. Malovic I, Sorensen KK, Elvevold KH, Nedredal GI, Paulsen S, Erofeev AV, Smedsrod BH, McCourt PA. The mannose receptor on murine liver sinusoidal endothelial cells is the main denatured collagen clearance receptor. Hepatology 45: 1454–1461, 2007.[CrossRef][Medline]
  41. Marciniak E, Gockerman JP. Heparin-induced decrease in circulating antithrombin-III. Lancet 2: 581–584, 1977.[Web of Science][Medline]
  42. Markwell MA. A new solid-state reagent to iodinate proteins. I. Conditions for the efficient labeling of antiserum. Anal Biochem 125: 427–432, 1982.[CrossRef][Web of Science][Medline]
  43. McCourt PA, Hansen B, Svistuonov D, Johansson S, Longati P, Schledzewski K, Kzhyshkowska J, Goerdt S, Smedsrod B. The liver sinusoidal endothelial cell hyaluronan receptor and its homolog, stabilin-1. Their roles (known and unknown) in endocytosis. Comp Hepatol 3, Suppl 1: S24, 2004.[CrossRef][Medline]
  44. McCourt PA, Smedsrod BH, Melkko J, Johansson S. Characterization of a hyaluronan receptor on rat sinusoidal liver endothelial cells and its functional relationship to scavenger receptors. Hepatology 30: 1276–1286, 1999.[CrossRef][Web of Science][Medline]
  45. Mego JL, Bertini F, McQueen JD. The use of formaldehyde-treated 131I-albumin in the study of digestive vacuoles and some properties of these particles from mouse liver. J Cell Biol 32: 699–707, 1967.[Abstract/Free Full Text]
  46. Melkko J, Hellevik T, Risteli L, Risteli J, Smedsrod B. Clearance of NH2-terminal propeptides of types I and III procollagen is a physiological function of the scavenger receptor in liver endothelial cells. J Exp Med 179: 405–412, 1994.[Abstract/Free Full Text]
  47. Monkhouse FC. Physiological factors concerned with the removal of injected heparin from the circulating blood. Am J Physiol 178: 223–228, 1954.[Free Full Text]
  48. Nakamura T, Yuasa H, Watanabe J. Uptake of fractionated heparin by two types of scavenger receptors in isolated rat Kupffer cells. Biol Pharm Bull 23: 743–747, 2000.[Medline]
  49. Oldberg A, Heldin CH, Wasteson A, Busch C, Hook M. Characterization of a platelet endoglycosidase degrading heparin-like polysaccharides. Biochemistry 19: 5755–5762, 1980.[CrossRef][Web of Science][Medline]
  50. Pertoft H, Smedsrod B. Separation and characterization of liver cells. In: Cell Separation: Methods and Selected Applications (vol. 4), edited by Pretlow TG and Pretlow TP. New York: Academic, 1987, p. 1–24.
  51. Praaning-van Dalen DP, Brouwer A, Knook DL. Clearance capacity of rat liver Kupffer, endothelial, and parenchymal cells. Gastroenterology 81: 1036–1044, 1981.[Medline]
  52. Rabinowitz S, Horstmann H, Gordon S, Griffiths G. Immunocytochemical characterization of the endocytic and phagolysosomal compartments in peritoneal macrophages. J Cell Biol 116: 95–112, 1992.[Abstract/Free Full Text]
  53. Selye H, Tuchweber B. Mast cell products and tissue calcification. Q J Exp Physiol Cogn Med Sci 50: 196–202, 1965.[Abstract/Free Full Text]
  54. Seternes T, Sorensen K, Smedsrod B. Scavenger endothelial cells of vertebrates: a nonperipheral leukocyte system for high-capacity elimination of waste macromolecules. Proc Natl Acad Sci USA 99: 7594–7597, 2002.[Abstract/Free Full Text]
  55. Slot JW, Geuze HJ. A new method of preparing gold probes for multiple-labeling cytochemistry. Eur J Cell Biol 38: 87–93, 1985.[Web of Science][Medline]
  56. Slot JW, Geuze HJ. Sizing of protein A-colloidal gold probes for immunoelectron microscopy. J Cell Biol 90: 533–536, 1981.[Abstract/Free Full Text]
  57. Slot JW, Geuze HJ, Gigengack S, Lienhard GE, James DE. Immuno-localization of the insulin regulatable glucose transporter in brown adipose tissue of the rat. J Cell Biol 113: 123–135, 1991.[Abstract/Free Full Text]
  58. Smedsrod B. Clearance function of scavenger endothelial cells. Comp Hepatol 3, Suppl 1: S22, 2004.[CrossRef][Medline]
  59. Smedsrod B, Einarsson M. Clearance of tissue plasminogen activator by mannose and galactose receptors in the liver. Thromb Haemost 63: 60–66, 1990.[Web of Science][Medline]
  60. Smedsrod B, Kjellen L, Pertoft H. Endocytosis and degradation of chondroitin sulphate by liver endothelial cells. Biochem J 229: 63–71, 1985.[Medline]
  61. Smedsrod B, Melkko J, Risteli L, Risteli J. Circulating C-terminal propeptide of type I procollagen is cleared mainly via the mannose receptor in liver endothelial cells. Biochem J 271: 345–350, 1990.[Web of Science][Medline]
  62. Smedsrod B, Pertoft H. Preparation of pure hepatocytes and reticuloendothelial cells in high yield from a single rat liver by means of Percoll centrifugation and selective adherence. J Leukoc Biol 38: 213–230, 1985.[Abstract]
  63. Smedsrod B, Pertoft H, Eggertsen G, Sundstrom C. Functional and morphological characterization of cultures of Kupffer cells and liver endothelial cells prepared by means of density separation in Percoll, and selective substrate adherence. Cell Tissue Res 241: 639–649, 1985.[Web of Science][Medline]
  64. Smedsrod B, Pertoft H, Gustafson S, Laurent TC. Scavenger functions of the liver endothelial cell. Biochem J 266: 313–327, 1990.[Web of Science][Medline]
  65. Stehle G, Friedrich EA, Sinn H, Wunder A, Harenberg J, Dempfle CE, Maier-Borst W, Heene DL. Hepatic uptake of a modified low molecular weight heparin in rats. J Clin Invest 90: 2110–2116, 1992.[Medline]
  66. Takata K, Horiuchi S, Araki N, Shiga M, Saitoh M, Morino Y. Endocytic uptake of nonenzymatically glycosylated proteins is mediated by a scavenger receptor for aldehyde-modified proteins. J Biol Chem 263: 14819–14825, 1988.[Abstract/Free Full Text]
  67. Teien AN. Heparin elimination in patients with liver cirrhosis. Thromb Haemost 38: 701–706, 1977.[Web of Science][Medline]
  68. Theroux P, Waters D, Lam J, Juneau M, McCans J. Reactivation of unstable angina after the discontinuation of heparin. N Engl J Med 327: 141–145, 1992.[Abstract]
  69. Valentin S, Larnkjer A, Ostergaard P, Nielsen JI, Nordfang O. Characterization of the binding between tissue factor pathway inhibitor and glycosaminoglycans. Thromb Res 75: 173–183, 1994.[CrossRef][Web of Science][Medline]
  70. Van Berkel TJ, Nagelkerke JF, Harkes L, Kruijt JK. Processing of acetylated human low-density lipoprotein by parenchymal and non-parenchymal liver cells. Involvement of calmodulin? Biochem J 208: 493–503, 1982.[Medline]
  71. Van Berkel TJ, Van Velzen A, Kruijt JK, Suzuki H, Kodama T. Uptake and catabolism of modified LDL in scavenger-receptor class A type I/II knock-out mice. Biochem J 331: 29–35, 1998.[Web of Science][Medline]
  72. Vannucchi S, Pasquali F, Chiarugi V, Ruggiero M. Internalization and metabolism of endogenous heparin by cultured endothelial cells. Biochem Biophys Res Commun 140: 294–301, 1986.[CrossRef][Web of Science][Medline]
  73. Watanabe J, Haba M, Muranishi H, Yuasa H. Dose-dependent uptake of radioactivity by liver parenchymal and non-parenchymal cells after intravenous administration of fractionated 3H-heparin to rats. Biol Pharm Bull 16: 1031–1034, 1993.[Medline]
  74. Watanabe J, Haba M, Yuasa H. Uptake of fractionated 3H-heparin by isolated rat Kupffer cells. Pharm Res 12: 1092–1095, 1995.[CrossRef][Medline]
  75. Watanabe J, Hori K, Iwamoto K, Ozeki S. Disposition of tritium-labeled heparin in rats. J Pharmacobio-Dyn 5: 627–637, 1982.[Medline]
  76. Watanabe J, Urano K, Muranishi H, Haba M, Yuasa H. Uptake of low molecular weight fractionated [3H]heparin by rat hepatocytes in the primary culture. Biol Pharm Bull 18: 443–446, 1995.[Web of Science][Medline]
  77. Young E, Venner T, Ribau J, Shaughnessy S, Hirsh J, Podor TJ. The binding of unfractionated heparin and low molecular weight heparin to thrombin-activated human endothelial cells. Thromb Res 96: 373–381, 1999.[CrossRef][Medline]
  78. Zhou B, Weigel JA, Fauss L, Weigel PH. Identification of the hyaluronan receptor for endocytosis (HARE). J Biol Chem 275: 37733–37741, 2000.[Abstract/Free Full Text]



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