The new anti-actin agent dihydrohalichondramide reveals fenestrae-forming centers in hepatic endothelial cells
© Braet et al; licensee BioMed Central Ltd. 2002
Received: 05 December 2001
Accepted: 21 March 2002
Published: 21 March 2002
Liver sinusoidal endothelial cells (LSECs) react to different anti-actin agents by increasing their number of fenestrae. A new structure related to fenestrae formation could be observed when LSECs were treated with misakinolide. In this study, we investigated the effects of two new actin-binding agents on fenestrae dynamics. High-resolution microscopy, including immunocytochemistry and a combination of fluorescence- and scanning electron microscopy was applied.
Halichondramide and dihydrohalichondramide disrupt microfilaments within 10 minutes and double the number of fenestrae in 30 minutes. Dihydrohalichondramide induces fenestrae-forming centers, whereas halichondramide only revealed fenestrae-forming centers without attached rows of fenestrae with increasing diameter. Correlative microscopy showed the absence of actin filaments (F-actin) in sieve plates and fenestrae-forming centers. Comparable experiments on umbilical vein endothelial cells and bone marrow sinusoidal endothelial cells revealed cell contraction without the appearance of fenestrae or fenestrae-forming centers.
(I) A comparison of all anti-actin agents tested so far, revealed that the only activity that misakinolide and dihydrohalichondramide have in common is their barbed end capping activity; (II) this activity seems to slow down the process of fenestrae formation to such extent that it becomes possible to resolve fenestrae-forming centers; (III) fenestrae formation resulting from microfilament disruption is probably unique to LSECs.
Liver sinusoidal endothelial cells (LSECs) differ from other endothelial cells. They possess open fenestrae that are grouped in sieve plates and lack a basal lamina . Fenestrae measure about 150 nm and occupy 6–8% of the endothelial surface (porosity) [2, 3]. The endothelial filter characteristics determine the exchange between the blood and the hepatocytes, and are affecting the hepatic metabolism of lipoproteins including cholesterol and vitamin A . Structural integrity of the fenestrated sinusoidal liver endothelium is believed to be essential for the maintenance of a normal exchange of fluids, solutes, particles and metabolites between the sinusoidal blood and hepatocytes. In the past, numerous publications appeared about the role of these dynamic structures under various physiological and pathological situations . Their role and involvement in the regenerating liver after partial hepatectomy , shear stress , liposome-mediated transport , liver cancer , injury by free radicals  and chronic alcohol abuse , resulting in alcoholism-associated hyperlipoproteinemia  have been explored.
In response to external signals, dynamic changes in fenestrae diameter and number occur and involve the dynamics of the actin cytoskeleton . Discoveries in the past decade have revealed that an actomyosin-driven machinery contributes to the regulation of fenestrae diameter and this under the control of intracellular calcium levels [14–16]. Detergent-extracted whole mounts of LSECs showed that fenestrae are delineated by a fenestrae-associated cytoskeleton ring (FACR), which changes in diameter and thickness after different treatments. These rings therefore seem to act as a supporting lattice and "muscle" around fenestrae . Furthermore, the actin cytoskeleton of LSECs has been shown to participate in cellular processes such as chemical- and cold-induced cell injury during liver transplantation [18, 19].
The recent availability of a battery of new actin binding drugs that affect the polymerization of actin by different mechanisms, provide a powerful tool to dissect the dynamics and functions of the actin cytoskeleton in various cell types . Previously it has been demonstrated that the treatment of LSECs with cytochalasin B , latrunculin A , swinholide A, misakinolide or jasplakinolide , induce an increase in the number of fenestrae. Only after treating LSECs with misakinolide, it was possible to resolve the process of fenestrae formation and to identify a new structure involved in the process of fenestrae formation . This illustrates the importance of the use of different anti-actin drugs to study the dynamic cellular processes that depend on the integrity and function of actin.
The present study endeavours to elucidate the dynamic process of de novo formation of LSEC fenestrae by correlating specific changes in actin organization with fenestrae. We used two novel actin-binding compounds derived from marine organisms, halichondramide (HALI) and dihydrohalichondramide (di-h-HALI), that belong to a large group of structurally related marine macrolides . Experiments on various cell lines show that HALI induces almost complete depletion of F-actin within minutes, illustrating the F-actin severing and monomer sequestering activities. Di-h-HALI, which differs from HALI only in having a single bond in the macrolide at positions 4–5 instead of a double bond, altered the filamentous F-actin distribution into large F-actin aggregates, and was found to possess strong barbed-end capping and weak severing activities . Both agents possess biochemical similarities to the previous tested anti-actin drugs swinholide and misakinolide which have severing and barbed-end capping activities, respectively [20, 23].
We report here that HALI and di-h-HALI: (I) disrupts actin organization in LSECs in a distinctive manner; (II) significantly increase the number of fenestrae; (III) that di-h-HALI elicits fenestrae-forming centers (FFCs) from which nascent fenestrae are fanning out; and (IV) for reasons of comparison, we also examined the effect of HALI and di-h-HALI on cultured human umbilical vein endothelial cells (HUVECs) and bone marrow sinusoidal endothelial cells (BECs STR-4). It was reported earlier that various treatments could induce fenestrae in HUVECs [24, 25]; whereas sinusoidal endothelial cells derived from the bone marrow posses the capacity to form transendothelial openings in vivo (i.e., transendothelial channels and diaphragmed fenestrae) while these structures are lost or greatly reduced in number in vitro [26, 27]. Therefore, the effect of HALI and di-h-HALI on BECs STR-4 was investigated to address the question whether these microfilament-disrupting drugs could induce fenestrae in this cell line derived from another sinusoidal source.
Lower concentrations of both compounds (25 and 50 nM) resulted in partial loss of F-actin bundles that were present even after 120 minutes incubation. Exposure to higher concentrations of HALI or di-h-HALI (200 nM) resulted in a decreased viability as assessed by the percentage of cells stained by propidium iodide: from 97% (control) to 59% and 65% respectively.
Scanning electron microscopy
While the maximum effect of the two drugs on actin organization was reached after 10 minutes, the maximum effect of the drugs on the number of fenestrae was reached at 120 minutes. Figure 2C depicts a di-h-HALI-treated LSEC after 120 minutes exposure to 100 nM di-h-HALI. The same SEM morphology was also seen in HALI-treated cells. Fenestrae were no longer clustered in sieve plates embedded in unfenestrated areas of cytoplasm (Fig. 2A), but treated cells contained abundant numbers of fenestrae interchanged with long and thin cytoplasmic arms, extending from the nucleus (Fig. 2C). Inside the fenestrated cytoplasm, the presence of small cytoplasmic unfenestrated areas could be observed. However, these areas were devoid of rows of very small fenestrae.
Fenestrae diameter (nm) after treatment with HALI or di-h-HALI obtained by scanning electron microscopy
Fenestrae diameter (nm)
Correlative fluorescence- and scanning electron microscopy
Transmission electron microscopy
Transversely cut sections revealed no additional structural information (data not shown).
Disassembly of filamentous actin in LSECs with the new microfilament-disrupting drug di-h-HALI enabled us to visualize FFCs (Figs. 2, 7, 9), most probably involved in the process of fenestrae formation, as demonstrated recently with the actin inhibitor misakinolide [20, 23]. Exposure of LSECs by other actin-perturbing agents including cytochalasin B , latrunculin A , swinholide A, jasplakinolide , and HALI (this paper) also produces a rapid increase in the number of fenestrae and the appearance of small unfenestrated areas which apparently represent inactive FFCs , indicating that actin disruption per se is sufficient to induce an increase in the number of fenestrae. However, the fact that only misakinolide and di-h-HALI resolved FFCs in the process of fenestrae formation indicate that specific alterations of the actin system are necessary to unmask active FFCs. Moreover, if our previous observations on FFCs as revealed by using misakinolide was an artefact of the drug, then it is most unlikely that di-h-HALI has the same side effect. The biochemical property that misakinolide and di-h-HALI have in common is their barbed end capping activity [20, 30], however misakinolide also forms actin dimers, whereas di-h-HALI possesses weak F-actin severing activity. In both cases an increased number of fenestrae (Fig. 3) and FFCs (Fig. 2, 7, 9)  could be observed, but misakinolide increases the number of fenestrae more rapidly than di-h-HALI (Fig. 3) , while di-h-HALI revealed approximately 40% more FFCs per squared micrometer (our unpublished data). While these differences may reflect distinct effects of these two agents on actin; it is possible that they also exert indirect effects on actin binding proteins as is the case with latrunculin . In contrast to the other anti-actin drugs that we tested, the specific alterations that misakinolide and di-h-HALI induce in the state of actin organization either by their barbed end capping activity or by indirect effects on the actin cytoskeleton appear to slow down the process of fenestrae formation to such an extent that it becomes possible to visualize active FFCs. In addition, beside these actin-related effects, possible membrane-associated interactions of di-h-HALI or misakinolide  cannot be excluded and may promote or inhibit the fusion/fission process of the cell membrane during fenestrae formation and as a consequence retard this process in such a way that FFCs appear with connected fenestrae rows. Intrinstingly, it has been recently reported that cytochalasin D facilitates apical membrane invagination and promotes exocytosis in pancreatic acinar cells ; whereas cytochalasin B inhibits membrane invagination recovery in neurons .
Endothelial cells of large vessels, which normally do not have fenestrae, have the ability to form fenestrae within minutes [34, 35]. This indicates that the process of fenestrae formation probably does not involve de novo synthesis of proteins, but rather a reorganization of preexisting cellular components. Indeed, specialized structures involved in the formation of diaphragmed fenestrae in the capillaries of the exocrine pancreas , adrenal cortex , kidney glomerulus  and tumor micro vascular endothelium  have been reported. It is presumed that peristomal rings of cholesterol, knob-like structures, vesiculo-vacuolar organelles and the endothelial pocket may represent important contributors to the formation of diaphragmed fenestrae. Therefore, it is conceivable that fenestrae increase, whether diaphragmed or not, is depending on pre-existing structures which promote fenestrae formation. Although these specialized structures may have an important role in the formation of diaphragmed fenestrae, their role in LSECs is less certain, primarily because LSEC fenestrae lack a diaphragm, are exceptionally abundant, and differ structurally from fenestrae in other blood vessels . The only fenestrae-related structure that LSECs and diaphragmed endothelial cells have in common in the complex process of fenestrae formation is the peristomal ring of sterols surrounding a fenestra . Nonetheless, our observations on the effect of di-h-HALI on large vessel endothelial cells and bone marrow sinusoidal endothelial cells (Fig. 5), demonstrated once more that the increase in the number of fenestrae and the appearance of FFCs by actin-disruption is probably a unique process for the hepatic sinusoidal endothelium. It has been reported that phorbol myristate acetate (PMA)  and vascular endothelial growth factor (VEGF) [25, 40] could induce diaphraghmed fenestrae in HUVECs. However, De Zanger et al.  showed that LSECs are insensible to PMA with regard to fenestrae induction. In addition, Krause and collaborators  recently noticed that the increase in the number of fenestrae with time in five days old LSECs cultures is independent of the presence of VEGF. Taking together, these observations clearly illustrate that LSECs respond in a different way to inducers of diaphragmed fenestrae, indicating once more that the machinery for the formation of diaphragmed versus non-diaphragmed fenestrae probably differs. Recent data are accumulating to show that VEGF-induced diaphragmed fenestrae are derived from fused caveolae . A recent study postulates that the same mechanism is used for the formation of LSEC-fenestrae [43, 44]. Evidence at the ultrastructural or molecular level about a possible relation between FFCs and interconnected caveolae is absent.
Fusion of two opposing cell membranes to form fenestrae probably requires the presence of unique compositional membrane microdomains and a cell membrane-associated cytoskeletal structure. Several theories have been used to model the possible mechanisms of membrane fusion for LSECs and other cell types. In general, the process leading to membrane fusion and fission is subdivided into different steps, i.e.: adhesion-dehydration; disappearance of the hydration barrier; contact between phospholipid bilayers, and; molecular rearrangement, resulting in pore formation [45, 46]. As for LSECs, the first step corresponds to the formation of intramembrane protein-free zones , while the appearance of peristomal rings of sterols probably corresponds with the final step . It seems reasonable to assume that these events take place in the rim of FFCs, and that these microdomains contain molecules to pull the bilayers of the cell membranes together at the edge of FFCs. Therefore, to define the FFC more precisely, we applied ultrathin sectioning and showed that filamentous structures of unknown nature are closely associated with these microdomains (Fig. 9C, 9D). In active FFCs, these filaments seem to serve as a guidance for the emerging nascent fenestrae. However, due to technical limitations it is impossible to get a nice plane overview picture of these filamentous structures on EM-sections. These limitations include the extremely small portion of the total cell volume that is included in a section, and the tendency of fine and thin structures cut obliquely or in cross section to disappear visually into the cytoplasmic ground substance. Together, these conditions make it almost impossible to correlate microscopical fluorescence data (Fig. 1B, 1C), with morphologically identified supramolecular structures in TEM (Fig. 9C, 9D). Therefore, in order to clarify the changes in actin organization that underlie the process of fenestrae formation correlative fluorescence- and SEM studies on the same cells was performed (vide infra). Surprisingly, examination of sectioned FFCs did not reveal additional structures regarding the architecture of this singular structure (Fig. 9C, 9D). Detailed investigation showed only a finely granular pattern of intermediate electron density. Although the molecular composition of FFCs remains unknown, F-actin was clearly found to be absent (Fig. 6I, 6L). Steffan et al.  postulated based on their in vitro and in situ studies with cytochalasin B that these pore-free microdomains may constitute an anchorage site for cytoskeletal elements. Our TEM sections (Fig. 9C, 9D) and correlative images (Fig. 6) support this statement but show that these cytoskeletal elements do not correspond with filamentous actin. In contrast, these F-actin patches clearly match with the fine globular topographic elevations present on the thin nonfenestrated cytoplasmic arms, and may represent anchor sites for linking F-actin with the plasma membrane .
Taking previous [20, 21, 23, 29, 47] and present findings together, we propose as an explanation for the events described that FFCs are anchored in the perinuclear area by the actin cytoskeleton where they cannot be resolved by electron microscopy due to their complex multi-fold organization. Disorganization of the filamentous actin cytoskeleton results in a centrifugally-like translocation of the FFCs towards the attenuated peripheral cytoplasm. Flattening of the FFCs occurs at the end of this movement and results in the appearance of FFCs with connected rows of fenestrae with increasing diameter. The spiraling rows may indicate that the translocated FFCs are rotating as they move into the peripheral region. The presence of a clear-cut FACR around every single fenestrae indicates that these centers already contain the necessary machinery and/or protein components for assembling the FACR around nascent fenestrae.
However, caution is required when interpreting this hypothesis, because electron microscopy provides only static information. Nevertheless, our data on the number of fenestrae rows connected to one FFC at different time points (Fig. 8), illustrates in one way a dynamic driven process at the level of the FFC. Recent attempts to confirm our hypothesis on the translocation of pre-existing FFC in real-time with atomic force microscopy failed [49, 50]. The unique low elastic modulus of living LSECs, together with the damaging tip-sample interactions constitute a problem to acquire sequential images of the process of fenestrae formation in real-time under the influence of cytoskeletal-disturbing agents.
A comparison of all anti-actin agents tested so far, revealed that the only activity that misakinolide and dihydrohalichondramide have in common is their barbed end capping activity; this activity seems to slow down the process of fenestrae formation to such extent that it becomes possible to resolve fenestrae-forming centers; fenestrae formation resulting from microfilament disruption is probably unique to LSECs.
Materials and methods
The method for the isolation of rat LSECs has been described in detail earlier . Briefly, the liver of a male Wistar rat (Center for Laboratory Animals, Leuven, Belgium – Rats received humane care in compliance with the institution's guidelines for the care and use of laboratory animals in research [accreditation number 99.1-212-3]) was perfused with collagenase A (Boehringer Mannheim, Catalogue Number 1088793, Belgium). After incubation of the fragmented tissue in the same solution, the resulting cell suspension was centrifuged at 100 × g for 5 minutes to remove the parenchymal cells. The supernatant, containing a mixture of sinusoidal liver cells, was then layered on top of a two step Percoll® gradient (25–50%) and centrifuged for 20 minutes at 900 × g. The intermediate zone, located between the two density layers was enriched in LSECs. LSEC purity was further enhanced by selective adherence of Kupffer cells and spreading of the LSECs on collagen. For SEM, LSECs were cultivated in 24-multiwell plates on collagen-coated thermanox cover slips. For whole mount TEM, LSECs were cultivated on collagen-coated nickel grids (300 mesh) instead of cover slips . Formvar (1%) supporting films on nickel grids (300 mesh) were used, coated with diluted collagen (10 μl of collagen-S stock solution [Boehringer Mannheim, Catalogue Number 1098292, Belgium], in 900 μl sterile water). Serum free LSEC culture medium consisted of RPMI-1640 with 2 mM L-glutamine, 100 U/ml penicillin, 100 μg/ml streptomycin and 10 ng/ml endothelial cell growth factor (Boehringer Mannheim, Catalogue Number 1074016, Germany). After 8 hours in culture, LSECs monolayers were extensively washed and subsequently used for experiments. The cultures were estimated to have greater than 95% purity, since less than 5% of the cells examined by EM were devoid of fenestrae.
HUVECs obtained from Clonetics Corporation (BioWhittaker, Catalogue Number CC-0216, New York) were grown to confluency in 24-multiwell plates with or without thermanox cover slips, using endothelial cell growth medium – 2 (BioWhittaker, Catalogue Number CC-3162, New York) supplemented with 2% foetal bovine serum (HyClon Labs, Logan, Utah). The cells were checked routinely for the presence of von Willebrand factor as described earlier .
The murine bone marrow sinusoidal endothelial cell line STR-4 (BEC STR-4) was established by transfecting primary endothelial cell cultures with SV40 . BECs STR-4 were maintained in culture in RPMI-1640 supplemented with 2 mM L-glutamine, 100 U/ml penicillin, 100 μg/ml streptomycin, MEM (Gibco Life Sciences, Catalogue Number 11120, Belgium) and 10% bovine serum (fetal Clone I, Hyclone, UT, USA). Cultures were regularly checked for von Willebrand factor positivity and the capacity to uptake LDL [53, 54].
Treatment of cells with HALI and di-h-HALI
LSECs and HUVECs were treated with 25, 50, 100 and 200 nM HALI or di-h-HALI for 10, 20, 30, 60 and 120 minutes. The compounds were dissolved in dimethyl sulfoxide (DMSO) and the DMSO concentration in the assays were in all cases ≤ 0.05% and this concentration of DMSO had no effect on the ultrastructure and viability of cells as determined by EM and Hoechst 33342 / propidium iodide staining. Control media also contained DMSO in the same amount as the treated cells. All experimental variables tested, including control cells, were fixed at the same end point, i.e. all cells underwent a two hours treatment with 0.05% DMSO and the compounds were added 10, 20, 30, 60 or 120 minutes prior to fixation [21–23]. After incubation and fixation, cells were prepared for fluorescence microscopy, and EM as described below.
In order to visualize simultaneously filamentous (F-actin) and globular (G-actin) actin, LSECs grown on glass cover slips were rinsed twice with phosphate-buffered saline (PBS) at pH 7.4, followed by fixation with freshly prepared 4% formaldehyde in PBS for 15 minutes at 21°C. After fixation, LSECs were submerged in absolute acetone for 3 minutes at -20°C. After this permeabilization, rhodamine-phalloidin (0.165 μM) and fluorescein-DNase I (0.3 μM) solution (Molecular Probes Inc., Eugene-Oregon, USA) was applied to LSECs for 20 minutes at 21°C. LSECs were subsequently washed 10 × with PBS and mounted on microscope slides in Vectashield containing DAPI (Vector Laboratories Inc., Burlingame, USA). As a control for the specificity of the staining reactions, LSECs were incubated first with 0.165 μM unlabeled phalloidin (Molecular Probes Inc., P-3457, Eugene-Oregon, USA) and 0.3 μM DNase I (Boehringer Mannheim S.A., Catalogue Number 104132, Germany) solution for 20 minutes at 21°C, before incubation with rhodamine-phalloidin and fluorescein-DNase I. No F- or G-actin staining was observed when control LSECs were incubated with unlabeled phalloidin or DNase I.
Cells were viewed and recorded with a Leica DM-IRBE inverted microscope, equipped with a Leica WILD MPS 48/52 – 35 mm camera. Elite Chrome 400 ASA slide film was used and slides were digitally scanned using a Minolta Dimâge Multi Scanner. The obtained images were transferred to Adobe Photoshop 5.5 software for color channel analysis and figure assembly. The magnification of the microscope was calibrated using fluoresbrite™ calibration grade microspheres, (Polylab BVBA, Ø 3.0 μm, Catalogue Number 18861, Belgium).
Scanning electron microscopy
Cells were rinsed twice with PBS and fixed with 2% glutaraldehyde in Na-cacodylate buffer (0.1 M and 0.1 M sucrose) at pH 7.4 for 12 hours. They were subsequently treated with filtered 1% tannic acid in 0.15 M Na-cacodylate at pH 7.4 for 1 hour and postfixed with 1% osmium tetroxide in 0.1 M Na-cacodylate at pH 7.4 for 1 hour. SEM samples were dehydrated in a graded ethanol series, dried with hexamethyldisilazane, and sputter coated with 10 nm of gold. The samples were examined with a Philips SEM 505 (Philips Eindhoven, The Netherlands) at an accelerating voltage of 30 KV .
For automatic image analysis, the SEM was regularly calibrated at a magnification of × 20,000, using a 28.800 lines/inch grating stub with the specimen in eucentric position. 30 images at a magnification of × 20,000 were taken in randomly selected fields of each experimental variable, each image containing a minimum of 10 fenestrae. Digital images with a low-noise content were obtained by using a large spot size (20 nm) and were processed subsequently and stored on a Masscomp 5520S computer, running under the RTU UNIX operating system, as previously described .
Correlative fluorescence- and scanning electron microscopy
Cells cultured on collagen-coated CELLocate-microgrid® glass cover slips (Eppendorf, Catalogue Number 5245 962.004-00, Hamburg, Germany) were treated with HALI or di-h-HALI as described and subsequently stained with rhodamine-phalloidin to visualize F-actin by fluorescence microscopy (see "Fluorescence microscopy" section). Photographs were then taken with the fluorescence microscope and photographed cells were located simultaneously by using the alphanumerically marked grids on the cover slip. Cover slips were recovered and transferred to 2% glutaraldehyde in Na-cacodylate buffer for 12 hours and further processed for scanning electron microscopy . Previously visualized cells in the fluorescence microscope were relocated with the aid of the alphanumerically marks on top of the cover slips and SEM-images of the corresponding cells were taken. Images of the same cells obtained from both microscopies were printed at an identical photographic end magnification , and were digitized using a Hewlett Packard ScanJet 3c scanner. The obtained images were transferred to Adobe Photoshop 5.5 software for color adjustment and figure assembly by using the replace color and duplicate layer/merge options.
Transmission electron microscopy
In order to visualize the cytoskeleton as a whole-mount for TEM , cells cultured on collagen-coated nickel grids were rinsed twice with PBS and slightly fixed for 1 minute with freshly prepared 4% formaldehyde in PBS at 21°C. Cells were subsequently extracted in cytoskeleton buffer consisting of 1 mM ethylene glycol bis [2-aminoethylether]-N,N,N',N' tetra-acetic acid, 100 mM piperazine-N,N'-bis [2-ethanesulfonic acid], 4% polyethylene glycol 6000 and 0.1% Triton X-100 in PBS at pH 6.9 for 1 minute at 21°C. After extraction, cells were processed as for SEM, but the tannin was omitted. Samples were further dehydrated and hexamethyldisilazane-dried. The specimens were examined in a Philips Tecnai 10 (Philips Eindhoven, The Netherlands) at an accelerating voltage of 100 kV.
For sectioning, cells cultured on cover slips were fixed and dehydrated in the same way as for SEM. After dehydration, samples were embedded in Epon and after hardening of the embedding medium, the cover slips were removed using liquid nitrogen. Sections of 60 nm under various angles were cut with a diamond knife, stained first with uranyl acetate, subsequently with lead citrate, and examined in a Philips Tecnai 10 at 80 kV as described .
All experiments were repeated five times. Statistical analysis was performed with the Mann-Whitney U test.
List of abbreviations used
liver sinusoidal endothelial cells
fenestrae-associated cytoskeleton ring
human umbilical vein endothelial cells
- BECs STR-4:
bone marrow sinusoidal endothelial cells STR-4
We thank Mrs Marijke Baekeland and Danielle Blijweert for expert technical assistance. The help of Mrs Chris Derom with excellent photographic assistance is very appreciated. The authors are also thankful to Prof. Dr. Michael Goligorsky and Dr. Jun Chen (Department of Medicine and Physiology – State University of New York at Stony Brook – USA) for the assistance in performing the experiments with the HUVEC monolayers. The authors thank Dr. M. Kobayashi (Laboratory of Pathology – Hokkaido University School of Medicine – Japan) for the generous gift of the STR-4 bone marrow sinusoidal endothelial cell line. This research was financially supported by the "Fund for Scientific Research-Flanders" (grant N° G000599 & G038000) and by the "Free University of Brussels" (I. Vanderschueren Price 2000 – Biomedicine [F.B.] & OZR230); and partially by the "National Oceanic and Atmospheric Administration Award" (NA46RG0090 NY Sea Grant Project R/XBP-5). E. Menu is an aspirant and F. Braet is a postdoctoral researcher of the "Fund for Scientific Research-Flanders (FWO-Fl)".
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