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An efficient, nonenzymatic method for isolation and culture of murine aortic endothelial cells and their response to inflammatory stimuli

Posted on: Thursday, 4 September 2003, 06:00 CDT

SUMMARY

Given the utility of murine models and the physiological and pathological significance of the aortic endothelium, we developed a simplified, nonenzymatic method for isolation and culture of murine aortic endothelial cells (MAECs). Aortic explants were initially cultured on fibronectin-coated plastic. Murine aortic endothelial cells migrated from the explants and proliferated. This expansion allowed for cultures to be established from the aortas of one or three mice. Murine aortic endothelial cells were then purified from expanded cultures by fluorescence-activated cell sorting for the uptake of 1,1'-dioctadecyl-3,3,3',3'-tetramethyl-indocarbocyanine perchlorate-labeled acetylated low-density lipoprotein. The majority of the cells in expanded cultures were as positive as human umbilical vein endothelial cells labeled in the same way. The most positive half of the labeled MAEC population was placed back in culture, and the cells formed "cobblestone" monolayers at confluence. Smooth muscle [alpha]-actin, which was present in aortic tissue and to a lesser extent in explant cultures before sorting, was not detected in selected MAECs. Western blotting and immunostaining also demonstrated the presence of the endothelial markers, platelet endothelial cell adhesion molecule-1, factor VIII- related antigen, and Bandeiraea simplicifolia lectin 1 binding. Murine aortic endothelial cells retained expected inflammatory functions: vascular cell adhesion molecule-1 protein was induced by bacterial endotoxin, and NO production was synergistically induced by the combination of endotoxin and Interferon-[gamma]. Our simple, efficient method will facilitate investigations of aortic endothelial cell function in vitro using murine models.

Key words: aorta; endothelium; mice; primary culture; endotoxin; interferon-[gamma].

INTRODUCTION

Endothelial cells line the inner surface of all blood vessels and play important roles in the regulation of vascular function throughout the body. Dysfunction of endothelium is important in many pathological conditions, including inflammation and atherosclerosis (Ross, 1999). Heterogeneity of endothelium can be dramatic because endothelial cells are specialized for different functions in different vascular sites (Kumar et al., 1987; Zetter, 1988). Endothelial cells from normal and diseased tissue may also differ markedly (Kumar et al., 1987; Gordon et al., 1991).

Although mice are generally inexpensive and amenable to transgenic modification, there are only a few methods reported for the culture of murine macrovascular endothelial cells. In contrast, there are many studies that have used macrovascular endothelial cultures from large animals such as cows, pigs, rabbits, and humans. The isolation of aortic endothelial cells from large animals typically begins with dissociation of cells from the vessel using enzymes such as collagenase and dispase. Enzymatic dissociation has not been as successful in rodents because of the resistance to digestion (Merrilees and Scott, 1981; McGuire and Orkin, 1987b). Enzymatic dissociation also promotes contamination of cultures with other vascular cells, such as smooth muscle cells (SMCs) and macrophages, which are liberated by enzymes, the risk increasing with duration oi digestion (Gimbrone et al., 1974; Gumkowski et al., 1987). Low cell viability is also a problem. These difficulties in mice have been addressed by starting with a large amount of tissue and growing cells in a selective medium (Gumkowski et al., 1987; Magee et al., 1994; Geiger et al., 1997) or selecting for the presence of endothelial markers (Auerbach et al., f982; Gumkowski et al., 1987; Kevil and Bullard, 2001).

A simple alternative to enzymatic digestion is explant culture. This method has been used for isolation of aortic endothelial cells from rats (McGuire and Orkin, 1987a, 1987b; Nicosia et al., 1994) and mice (Kondo et al., 1994; Suh et al., 1999). Aortic explants are typically cultured with the endothelial side of the vessel in contact with extracellular matrix coatings such as collagen, fibronectin, or matrigel, to enhance endothelial outgrowth and survival. These explants readily produce a large population of cells from a small amount of tissue through proliferation. The previously reported murine cultures were predominantly endothelial in composition (Kondo et al., f994; Suh et al., 1999). However, as with enzymatic dissociation, contamination with other cell types can occur (McGuire and Orkin, 1987a, 1987b; Nicosia et al., 1994; Suh et al., 1999). Smooth muscle cells and fibroblasts appear to be the major nonendothelial cell types that persist (Jaffe et al., 1973; Suh et al., 1999). To date, the selection of cells for endothelial markers, which has proven successful after enzymatic dissociation, has not been reported for marine aortic explant cultures.

In this study, we used fibronectin-coated plastic to establish aortic explant cultures starting with one or three mice. The cell population was expanded by two passages on plastic. Fluorescence- activated cell sorting (FACS) for uptake of 1,'-dioctadecyl- 3,8,3',3'-tetramethyl-indocarbocyanine perchlorate-labeled acetylated low-density lipoprotein (di-I-acetylated-LDL) was then used to purify endothelial cells from these subcultures (Voyta et al., 1984). Immunostaining and Western blotting for specific cell markers were used to demonstrate the cells' endothelial nature. The response of the cells to known inflammatory stimuli was measured to demonstrate their functional state. Our method is a simple, efficient, and cost-effective alternative to other methods for murine aortic endothelial cell culture.

MATERIALS AND METHODS

C57B1/6 mice, weighing 25-40 g each, were obtained from Jackson laboratories, Bar Harbor, Maine. NIH/3T3 cells were obtained from American Type Culture Collection, Rockville, Maryland. Human umbilical vein endothelial cells (HUVECs) were purchased from Clonetics, East Rulherford, New Jersey.

Sodium pentobarbital was obtained from Abbott Laboratories, North Chicago, Illinois. Dulbecco minimum essential medium (DMEM), F-12 nutrient mixture with L-glutamine (HAM), minimal essential medium lacking calcium (S-MEM), 0.25% trypsin-1 mM ethylenediaminetetraacetic acid (trypsin-EDTA), phosphate-buffered saline (PBS), penicillin-streptomycin solution (200 U penicillin and 200 [mu]g streptomycin per milliliter), and Tris-glycine gels were from Invitrogen Corporation, Grand Island, New York. Fetal bovine serum (FBS) was from Hyclone, Logan, Utah. Endothelial cell growth supplement (ECGS; bovine neural tissue extract), heparin, bovine serum albumin (BSA), Escherichia coli endotoxin serotype 0111:B4 (LPS), actinomycin D, cycloheximide, sulfanilamide, naphthalene- ethylenediamine dihydrochloride, M^sup c^-methyl-L-arginine (L- NMMA), aminoguanidine, fluorescein-conjugated Bandeiraea simplicifolia lectin I (BS), D-galactose, and Bradford reagent were from Sigma Chemical Co., St. Louis, Missouri. Escherichia coli- denved recombinant mouse Interferon-[gamma] (IFN-[gamma]) was from R&D systems, Minneapolis, Minnesota. Fibronectin was from Calbiochem, La Jolla, California, and di-I-acetylated-LDL was from Biomedical Technologies, Inc., Stoughton, Massachusetts. Triton X- 100 was from Pierce, Rockford, Illinois. Tris-base, sodium chloride, EDTA tetrasodium salt, sodium vanadate (ortho), sodium fluoride, and Tween 20 were from Fisher Scientific, Fair Lawn, New Jersey. Aprotinin, leupeptin, and pepstalin A were from Boehringer Mannheim Corporation, Indianapolis, Indiana. Chemiluminescence reagents were obtained from NEN Life Science Products, Inc., Boston, Massachusetts. Goat anti-human factor VIII-related antigen (factor VIIIra) antibody and fluorescein isothiocyanale (FITC)-conjugated rabbit anti-goat IgG were from ICN Biomedicals, Inc., Costa Mesa, California. Rat anti-mouse CD31 (platelet endothelial cell adhesion molecule-1, PECAM-1), rat anli-mouse CD106 (vascular cell adhesion molecule-1, VGAM-1), rat anti-mouse intercellular adhesion molecule- 1 (ICAM-), and rat anti-mouse integrin ssl polyclonal antibodies for immunostaining were obtained from Pharmingen, San Diego, California. Goat anti-mouse PECAM-1 and VCAM-1 antibodies for Western blotting were from Santa Cruz Biotechnology, Inc., Santa Cruz, California. Mouse anti-actin (smooth muscle) antibody was obtained from Research Diagnostics, Inc., Flanders, New Jersey. Rabbit polyclonal antibody to inducible nitric oxide synthase (iNOS) was obtained from Transduction Laboratories, Lexinglon, Kentucky. Cy3- or horseradish peroxidase (HRP)-conjugated goat anti-mouse, goat anti-rat, goal anti-rabbit, and rabbit anti-goat secondary antibodies were from Jackson ImmunoResearch Laboratories, Inc., West Grove, Pennsylvania.

Cell isolation. The thoracic aortas were removed, without stretching, from anesthetized mice (sodium pentobarbital, 300 mg/kg body weight, intraperitoneal). The vessels were rinsed two to three times and kept in endothelial cell culture medium (40% DMEM, 40% HAM, 20% FBS [heat inactivated, 30 min at 56[degrees] C], 30 [mu]g/ ml ECGS, 10 U/ml heparin, and penicillin-streptomycin). The periadventitial fat around the vessels was carefully cleaned under a dissecting microscope \using forceps and iris scissors. After opening the vessels longitudinally, each one was cut into four to six small pieces of approximately 1- to 2-mm^sup 2^ and placed in a 9.6-cm^sup 2^ fibroneetin-coated culture well. Coating of wells was accomplished, as described previously, by incubation with 2 ml fibronectin (50 [mu]g/ml PBS) for 2 h at 37[degrees] C. Dishes were then washed twice with PBS, air-dried, and stored at 4[degrees] C until use (Hoyt et al., 1996). After placing the pieces of aorta (endothelial cell side down) in the dish, a small amount of medium was added to keep the tissue moist without submerging them. This prevented the tissue from floating, allowing the pieces to remain in contact with the fibronectin-coated substratum. The explants were placed in an incubator at 37[degrees] C in 5% CO2 atmosphere.

Explants attached to the surface after 1-2 d in this minimal volume of medium. At this point, 0.5 ml of medium was carefully added so as to prevent dislodging the tissue. Cells began to migrate from the edges of explants between days 2 and 7, depending on the age of mice used. Usually, endothelial outgrowths were observed within 2-3 d when mice less than 8 wk of age were used. The medium in explant cultures was routinely changed every 2-3 d, and explants were removed and discarded when cells were still subconfluent (i.e., 70-80% of available area was occupied in 7-10 d using material from a single young mouse). Cells were then cultured for 2-3 d more until just confluent. Trypsin was used to detach cells for further subculture in normal 25-cm^sup 2^ plastic culture flasks without fibronectin. These cells grew to confluence within 7-14 d. Alternatively, cells from three mice cultured in separate 9.6- cm^sup 2^ fibronectin-coated wells were pooled in a single 25- cm^sup 2^ plastic culture flask for 1 d. In both cases, the first 25- cm^sup 2^ plastic culture was termed "passage 1."

Fluorescence-activated cell sorting. When cells reached confluence, they were detached from the 25-cm^sup 2^ flask with trypsin. Eighty percent of the cells were placed in a 75-cm^sup 2^ flask (passage 2), and 20% were placed in a 25-cm^sup 2^ flask. When the cells were approximately 80% confluent, 5 [mu]g di-I-acetylated- LDL/ml cell culture medium was added to the 75-cm^sup 2^ flask. The 25-cm^sup 2^ flask was incubated with medium only and was used as an autofluorescent blank for FACS. The following day, the cells were detached with trypsin and resuspended in FACS buffer (S-MEM lacking calcium-0.5% BSA-penicillin-streptomycin, filter sterilized) at a concentration of 1 x 10^sup 6^ cells/ml. The cells were kept in a tube on ice until they were sorted with a Beckman-Coulter Elite I ESP Flow Cytometer, using argon laser illumination (488 nm) and a 550-nm band pass emission filter for detection, essentially as described previously (Voyta et al., 1984). Human umbilical vein endothelial cells were labeled with and without the modified low- density lipoprotein (LDL) and sorted in the same way. Cells were collected in 50% medium-50% serum. The brightest half of the murine population was collected and placed in culture (passage 3). These murine aortic endothelial cells (MAECs) were subcultured by splitting at a ratio of 1:3, using untreated plastic culture flasks.

Cell characterization by immunostaining. The expression of factor VIIIra, PECAM-1, VCAM-1, ICAM-1, and integrin [beta]1 was determined. Cells were cultured in 16-well glass chamber slides and were allowed to grow overnight. The medium was removed, and the cells were rinsed with ice-cold PBS three times to remove any residual serum. Murine aortic endothelial cells were fixed with ice- cold acetone for 10 min. After three washes with ice-cold PBS, the cells were blocked with 10% rabbit serum (for factor VIIIra staining) or 10% goat serum (for PECAM-1, VCAM-1, ICAM-1, or integrin [beta]1 staining) at 37[degrees] C for 1 h. The cells were then incubated with different primary anti-bodies (goat anti-human factor VIIIra IgC, 20 [mu]g/ml in 10% rabbit serum; rat anti-mouse PECAM-1, 10 [mu]g/ml in 10% goat serum; rat anti-mouse VCAM-1, 10 [mu]g/ml in 10% goat serum; rat anti-mouse ICAM-1, 10 [mu]g/ml in 10% goat serum; or rat anti-mouse integrin [beta]1, 5 [mu]g/ml in 10% goat serum) overnight at 4[degrees] C. Control cells were incubated with preimmune goal serum in 10% rabbit serum (for factor VIIIra staining) or preimmune rat serum in 10% goat serum (for PECAM- 1, VCAM-1, ICAM-1, or integrin [beta]1 staining). The cells were then washed with PBS three times and incubated in either FITC- conjugated rabbit anti-goat IgG (7.5 [mu]g/ml for factor VIIIra staining) or Cy3-labeled goat anti-rat IgG (3.5 [mu]g/ml for PECAM- 1, VCAM-1, ICAM-1, or integrin [mu]1 staining) at 37[degrees] C for 1 h. The free secondary antibody was washed out with PBS after the incubation, and the cells were detected under a fluorescence microscope. NIH/3T3 fibroblasts, a nonendothelial negative-control cell type, were labeled in the same way. Cells were also fixed with 1% formaldehyde in medium at 37[degrees] C for 10 min, rinsed with PBS, and stained with fluorescein-conjugated BS1, in the absence and in the presence of 500 mM D-galactose to displace lectin specifically bound to endothelial cells (Sahagun et al., 1989).

Western, blotting. Western blotting was used to assess the endothelial cell marker, PECAM, and SMC/fibromyocyte-specific [alpha]-actin. Induction of VCAM-1 and iNOS by proinflammatory agents was also measured.

FIG. 1. Mouse aortic- explant and endothelial cells viewed by phase-contrast microscopy. Endothelial cells started to migrate from the edges of explants in 2-3 d. (A) An edge of an explant, lower left corner (arrow), and monolayer of cells with cobblestone appearance at day 7. This typical endothelial cell morphology was evident from passage 3, the first passage after fluorescence- activated cell sorting (B) through passage 20 (C) and beyond (data not shown). Bar in panel A, 100 [mu]m.

Western blotting was performed on extracts of the purified MAECs and on mouse aortic tissue. Murine aortic endothelial cells or tissue extracts were prepared in cell lysis buffer (1% Triton X- 100, 50 mM Tris, pH 7.5, 150 mM NaCl, 5 mM EDTA, 0.5 mM Na^sub 3^VO^sub 4^, 50 mM NaF, 10 [mu]g/ml aprotinin, 10 [mu]g/ml leupeptin, and 1 [mu]g//ml pepstatin A). The lysates were sonicated, and the proteins were heated at 95[degrees] C for 10 min. Ten micrograms of protein from cell or tissue extracts was separated on 4-20% Tris-glycine gel. After electrophoresis, the proteins were transferred onto nitrocellulose membranes. After transfer, the membranes were blocked with 3% nonfat milk in Tween 20-Tris- buffered saline buffer (0.1% Tween 20; 10 mM Tris, pH 7.5; 150 mM NaCl) and incubated with mouse anti-smooth muscle [alpha]-actin antibody (1 [mu]g/ml) for 2 h at room temperature and then with goat anti-mouse HRP-coupled secondary antibody. The blots were developed with an enhanced chemiluminescence system. The chemiluminescent signals were captured on X-ray film, scanned, and quantified by digital image analysis (Image J 1.22d, NIH). The integrated total signal intensify for bands was calculated from this densitometric information.

LPS-stimulated VCAM-1 and iNOS expression was assessed in MAECs pretreated with LPS at 100 [mu]g/ml for 2 h at 37[degrees] C. Murine aortic endothelial cells were also treated with 0-100 [mu]g LPS/ml for 0.5-24 h, in the absence or presence of IFN-[gamma] (1-20 ng/ ml). In some experiments, the cells were preincubated with either actinomycin D (1 [mu]g/ml) or cycloheximide (30 [mu]g/ml) for 16 h before the LPS treatment. Whole-cell extracts and Western blots were then prepared as described above. The blots were incubated for 1 h with goat anti-mouse monoclonal anti-VCAM-1 antibody and rabbit anti- goal HRP-coupled secondary antibody or rabbit anti-iNOS antibody and goat anti-rabbi t-HRP antibody.

LPS and IFN-[gamma]-stimulated NO production. The concentration of nitrite in cell culture supernatant, which reflects cumulative NO production, was measured by the Greiss reaction (Finder et al., 1995). Murine aortic endothelial cells were stimulated with 0-100 [mu]g/ml LPS for 0-24 h, with or without IFN-[gamma] (1-20 ng/ml). In some experiments, the cells were coincubated with L-NMMA (0.1 [mu]M to 1 mM) or aminoguanidine (0.1 [mu]M to 1 mM). Cultured supernatant (50 [mu]l) was sampled and immediately mixed with 50 [mu]l of Greiss reagent (1% sulfanilamide, 0.1% naphthalene- ethylenediamine dihydrochloride in 5% H^sub 3^PO^sub 4^). After incubation for 15 min at room temperature, the samples were read at 550 nm using a microplate spectrophotometer (Molecular Devices, Sunnyvale, CA). Solutions of sodium nitrite diluted in culture media served as the standard. A standard curve was made for every experiment carried out. The levels of nitrite were further normalized to the protein level in each sample. Protein was measured after washing cells with PBS three times and extraction with 1% Triton X-100. The samples were centrifuged at 3000 rpm for 10 min. Protein levels were then determined by the method of Bradford (Bradford, 1976).

FIG. 2. Representative fluorescence-activated cell sorting of cells incubated with 1,1'-dioetadecyl-3,3,3',3'-tetramethyl- indocarbocyanine perchlorate-labeled acetylated low-density lipoprotein (di-I-acetylated-LDL). The horizontal axis represents the logarithm of the relative fluorescence. The vertical axis represents the number of cells at each fluorescence level. (Panel A) Human umbilical vein endothelial cells (HUVECs) were incubated without the marker to define the autofluorescence of the blank, indicated by horizontal line segment B in each panel. (Panel B) Human umbilical vein endothelial cells, preincubated with di-I- acetylated-LDL (5 [mu]g/ml, 24 h), were fluorescent (marked by horizontal line segment C in the panel). Mouse c\ells were also preincubated for 24 h in the absence (panel C) and presence (panel D) of di-I-acetylated-LDL. Eighty-four percent of the mouse cells in this example were in the fluorescence intensity range defined by HUVEC staining in panel B (horizontal line segment C). The upper half of the mouse cell population was recovered for further culture and characterization as murine aortic endothelial cells.

FIG. 3. Western blotting for platelet endothelial cell adhesion molecule-1 in murine aortic endothelial and NIH/3T3 cell extracts. Ten micrograms of protein was subjected to electrophoresis and blotting as described in Materials and Methods. The position of molecular weight markers is indicated at the left. Murine aortic endothelial cells (passage 16) and 3T3 samples are indicated at the top.

FIG. 4. Western blotting for smooth muscle-specific [alpha]- actin in extracts of murine aortic endothelial cells and aortic tissue. Murine aortic endothelial cells from primary culture or subculture (passage 3 through 20, P3-P20) had no observable [alpha]- actin. Cells extracted before fluorescence-activated cell sorting for di-I-acetylated LDL uptake (US) and aortic tissue had detectable levels, consistent with the significant presence of smooth muscle cells.

Statistical testing. All data were evaluated by analysis of variance, with Bonferroni correction for multiple comparisons (Snedecor and Cochran, 1980). A P value less than 0.05 was considered significant.

RESULTS

Phase-contrast microscopy. During the first 2-3 d after the aortic explants attached to the 9.6-cm^sup 2^ fibronectin-coated wells, small colonies of cells migrated from the edges, each containing two to five polygonal cells with large, round nuclei. Migration continued, and marked proliferation began at 5-7 d after isolation. Wells were confluent 10-14 d after starting the cultures (Fig. 1a). Subculturing a single well from a single mouse in the first 25-cm^sup 2^ plastic flask yielded a confluent, "cobblestone" monolayer, typical of macrovascular endothelial cells, within 7-14 d. Naturally, pooling three 9.6-cm^sup 2^ fibronectin-coated wells (i.e., three mice) immediately produced the confluent 25-cm^sup 2^ flask (passage 1), which was split the following d between a 75- cm^sup 2^ flask and a 25-cm^sup 2^ flask for FACS (passage 2). This pooling of samples eliminated the extra 7-14 d necessary for samples from individual mice. The total time from starting expiants until cells were ready for di-I-acetylated-LDL labeling at passage 2 was 14-21 d for the pooled cultures and 21-28 d for separate cultures from individual mice.

FIG. 5. Western blotting for vascular cell adhesion molecule-1 (VCAM-1) in Escherichia coli endotoxin serotype 0111:B4 (LPS)- treated murine aortic endothelial cells. (A) Ten micrograms of LPS per milliliter caused a time-dependent increase in VCAM-1 expression within 4 h. The induction was blocked by 16-h preincubation with either 1 [mu]g actinomycin D/ml (4/ActD) or 30 [mu]g cycloheximide/ ml (4/CHX) (** P < 0.01, *** P < 0.001 for comparison with time 0; +++ P < 0.001 for comparison with 10 [mu]g LPS/ml, 4-h treatment; n = 3). (B) The induction of VCAM-1 by LPS (open bars) was concentration dependent and lasted at least 12 h. Coincubation with 20 ng interferon-[gamma] (IFN-[gamma]) per milliliter (solid bars) enhanced the effect of LPS slightly but significantly (*** P < 0.001 for comparison with no LPS or IFN-[gamma] treatment; ## P < 0.01, P < 0.001 for comparison with LPS only; n = 3). Representative Western blots are shown, the lanes positioned over the corresponding bars in the graphs below them.

Cultures selected by FACS for di-I-acetylated-LDL uptake retained the typical endothelial cell morphology (Fig. 1b, passage 3). This appearance remained throughout subculture. The cells grew rapidly in the complete medium containing ECGS and heparin and survived normal cell freezing and liquid nitrogen storage procedures without morphological changes after thawing and continued culture (Fig. 1c).

Fluorescence-activated cell sorting. Endothelial cells and macrophages selectively Lake up and degrade acetylated LDL (Voyta et al., 1984). This properly distinguishes them from fibroblasts, SMCs, and pericytes, which are common contaminants in primary endothelial cultures. We incubated HUVECs and murine explant cells with the fluorescent derivative of acetylated LDL, di-I-acetylated-LDL. By comparison with the lowest fluorescence of di-T-acetylated-LDL- treated HUVECs, the large majority of primary murine cells were deemed positive (84% in the example in Fig. 2). As mentioned above, the brightest 50% of the murine cell populations were collected for further culture.

FIG. 6. NO production as reflectedd by nitrite in medium. (A) Kinetics of NO production in murine aortic endothelial cells treated with [mu]g LPS per milliliter in the absence (open bars) or presence of 20 ng interferon-[gamma] (IFN-[gamma]) per milliliter (solid bars). (*** P < 0.001 for comparison with lime 0; P < 0.001 for comparison with LPS only; n = 6.) (B) Effect of different concentrations of LPS and IFN-[gamma] (* P < 0.05, *** P < 0.001 for comparison with 0 [mu]g LPS/ml; n = 6).

Endothelial markers. Western blotting demonstrated the presence of PECAM in MAECs and its absence in the nonendothelial NIH/3T8 cells (Fig. 3). A large amount of smooth muscle [alpha]-actin, which is characteristic of SMCs and myofibroblasts of the vessel wall, was present in extracts of murine aortic tissue (Fig. 4). A lesser signal was detected in explant MAEC cultures before sorting (passage 2), suggesting the presence of some of these nonendothelial cell types. In contrast, the amount of smooth muscle [alpha]-actin was too low to be observed in the di-I-acetylated-LDL-positive MAECs, suggesting that FACS enriched the endothelial cell content of the cultures. The selected MAECs were also found to be positive for factor VIIIra, PECAM-1, BS1 binding, basal VCAM-1, ICAM-1, and integrin [beta]1 by staining and fluorescence microscopy (not shown).

Upregulalion of VCAM-1 by LPS. Cytokines and other inflammatory stimuli induce adhesion molecules on endothelial cells that allow leukocytes to recognize sites of inflammation and to transmigrate through the blood vessel wall (Osborn, 1990; Gerritsen et al., 1995). Vascular cell adhesion molecule-1 is one such adhesion molecule, and it was induced by LPS alone in MAECs (Fig. 5). There was a time-dependent increase in VCAM-1 levels in the first 4 h after treatment. Maximal levels were reached at 4-6 h and held for at least 12 h. Induction was dose dependent, and IFN-[gamma] enhanced the effect of LPS slightly. Actinomycin D (1 [mu]g/ml) and cycloheximide (30 [mu]g/ml) abolished the induction, but not basal expression, suggesting that induction was attributable to increased transcription and translation. The induction of VCAM-1 by LPS was also visible by immunoflourescence and microscopy (data not shown).

Upregulation of iNOS by LPS and IFN-[gamma]. NO production during inflammation regulates vascular tone and other vascular cell functions. Inflammatory stimuli and cytokines typically induce iNOS in endothelial cells. We measured nitrite levels, which reflect original NO, in the medium of cells challenged with LPS alone or in combination with IFN-[gamma] (Fig. 6). Whereas neither agent alone had any effect, the combination of LPS and IFN-[gamma] greatly increased nitrite levels. As expected, NO production was blocked by both L-NMMA, a nonspecific nitric oxide synthase (NOS) inhibitor (the concentration causing 50% inhibition [IC^sub 50^] = 43 [mu]M, Fig. 8a), and aminoguanidine, which is a more specific iNOS inhibitor (the concentration causing 50% inhibition [IC^sub 50^] = 180 [mu]M, Fig. 8b).

Production of NO was delayed until 10 h after treatment with LPS and IFN-[gamma] (Fig. 6a) but then continuously increased through 24 h. The synergistic increase depended on the concentrations of both LPS and IFN-[gamma] (Fig. 6b). Western blotting for iNOS protein demonstrated that LPS plus IFN-[gamma] induced the enzyme within 6 h. The protein level continued to increase up to 24 h after treatment (Fig. 7). In contrast to VCAM-1, the potentiation of induction of iNOS protein in MAECs treated with 5 [mu]g LPS/ml and 20 ng IFN-[gamma]/ml was 10- to 20-fold at 24 h with respect to cells treated with LPS alone (Fig. 7). The potentiation of VCAM-1 induction by IFN-[gamma] was only about twofold in MAECs treated the same way. The lower ratio is because of the induction of VCAM-1 by LPS alone (Fig. 5b).

DISCUSSION

Endothelial cells have been isolated from murine aortas after perfusion and digestion with collagenase or collagenase plus dispase (Cumkowski et al., 1987; Kevil and Bullard. 2001). In these studies, endothelial cells were subsequently enriched by FACS for di-I- acetylated-LDL uptake (Gumkowski et al., 1987) or for binding of FITC-conjugated BS1 (Kevil and Bullard, 2001). Potential enzyme- induced contamination with nonendothelial cell types was overcome by these selection procedures. Twelve mice were processed and pooled for each preparation in the latter study, and rapid and careful perfusion with collagenase and ligation of vessels for digestion were credited with maintenance of adequate cell viability (Kevil and Bullard, 2001). In this study, we also found that it was important to work rapidly when isolating and cleaning aortas to minimize the time that explants spent ex vivo before placement in culture medium at 37[degrees] C. Also, vessels were manipulated carefully to avoid injury to the endothelial lining.

FIG. 7. Western blotting for inducible nitric oxide synthase (iNOS) in LPS-treated murine aortic; endothelial cells. LPS alone (open bars) had no effect on iNOS protein levels. The combination of LPS and 20 ng IFN-[gamma] per milliliter synergistically induced iNOS protein, which was both time- and dose-dependen\t (*** P < 0.001 for comparison with no LPS or IFN-[gamma] treatment; P < 0.001 for comparison with LPS only; n = 3). Representative Western blots are shown, the lanes positioned over the corresponding bars in the graphs below them.

Our method offers a flexible and convenient alternative to enzyme perfusion and digestion. Simple tissue harvesting and explant culture are performed on one d, and FACS is reserved for a later time alter expansion of the explant endothelial cells. Furthermore, it was found that single mice yielded primary explant cultures ready for FACS in 21-28 d. This could be useful if individual mice are required for investigation. If not, tissue from multiple animals can be pooled. Typically, pooling vessels from three mice produced cultures for FACS in 14-21 d. The explant method also minimizes the period during which cells are detached from a culture surface, which can induce endothelial cell apoptosis (Meredith et al., 1993). It may also be possible to culture endothelial cells from different segments of the aorta, which could be important for investigation of certain diseases that develop preferentially in different areas of the vessel (Iiyama et al., 1999).

Fibronectin was used as in other studies to promote adhesion, survival, and growth of primary endothelial cells (Meredith et al., 1993; Gerritsen et al., 1995; Hoyt et al., 1996; Hoyt et al., 1997). Fibronectin acts partly by associating with cellular [alpha]5[beta]1 integrin through arginine-glycine-aspartate sequences in its structure (Hynes, 1992). In pilot studies, we found that initial explants behaved similarly on fibronectin and the less expensive protein, gelatin, which also associates with [beta]1 integrins. Murine aortic endothelial cells did express [beta]1 integrins as expected. Matrix coatings were not required for continued culture of the cells.

The combination of ECGS and heparin can support long-term culture of endothelial cells. Endothelial cell growth supplement and heparin, respectively, stimulate endothelial cell replication and inhibit SMC proliferation (Castellot et al., 1982; Tan et al., 1989). Heparin also potentiates the mitogenic action of ECGS (Thornton et al., 1988; Tan et al., 1989). A high concentration of FBS (20%) was used in the present study to enhance outgrowth (McGuire and Orkin, 1987b). Using this medium, growth arrest has not been observed, even beyond 30 passages at a 1:3 split ratio. We have confined the present observations to cells under 20 passages and find that the reported characteristics and markers are retained (Figs. 1 and 4 for examples). They may persist for longer times.

Although several groups have used explant culture of endothelial cells from small animals, either technical difficulties or contamination with other cell types was observed (McGuire and Orkin, 1987a, 1987b; Nicosia et al., 1994). Smooth muscle cells and fibroblasts are potentially major sources of contamination of endothelial cell cultures (Jaffe et al., 1973). Indeed, smooth muscle [alpha]-actin was detected in the initial explant cultures in the present study (Fig. 4).

Di-I-acetylased LDL is taken up by endothelial cells and macrophages through the lipoprotein scavenger receptor (Voyta et al., 1984) but to a much lesser extent by aortic smooth muscle and fibroblasts (Stein and Stein 1980; Modzelewski et al., 1994). Fluorescence microscopy or flow cytometry for this marker has often been used to characterize endothelial cell cultures in general and to purify endothelial cells from enzymatically digested mouse aorta (Gumkowski et al., 1987). However, FACS for di-I-acetylated-LDL uptake by expanded murine aortic explant cultures has not been reported. The majority (>80%) of primary murine explant cells here were as positive for di-I-acetylated-LDL uptake as HUVECs analyzed at the same time. The brightest half of the murine cell populations was retained for culture as a conservative measure. It may be possible to lower this threshold for collection and increase the initial number of cells recovered. However, cell number is not a limiting factor when using cultures that have been expanded before labeling.

Alter sorting, the MAECs grew as a cohesive sheet of large polygonal cells, with oval, centrally located nuclei. Murine aortic endothelial cells were positive for the endothelial markers, PECAM- 1 (Fig. 3), factor VIIIra, and BS1 binding (Albelda et al., 1991; Modzelewski et al., 1994; Kevil and Bullard, 2001). Factor VIIIra- negative cells were not observed, suggesting that macrophages, which also ingest di-I-acetylated-LDL, were not present. The FACS- selected MAECs lacked detectable smooth muscle a-actin, suggesting minimal contamination with SMCs and fibroblasts (Fig. 4). Thus, FACS was effective in enriching cultures of MAECs, which appeared to remain free of contaminating cell types for an extended number of passages.

FIG. 8. Effect of nitric oxide synthase (NOS) inhibitors on LPS and IFN-[gamma]-induced NO production. (A) Murine aortic endothelial cells were coincubated with N^sup G^-methyl-L-arginine, a nonspecific NOS inhibitor. The production of NO by LPS and IFN- [gamma] was inhibited in a concentration-dependent manner (the concentration causing 50% inhibition [IC^sub 50^] = 43 [mu]M). (B) Murine aortic endothelial cells were coincubated with aminoguanidine, a specific inducible nitric oxide synthase inhibitor. The production of NO was also inhibited (IC^sub 50^ = 180 [mu]M) (* P < 0.05 and *** P < 0.001 for comparison with treatments in the absence of the inhibitors; n - 6).

We measured two typical responses to proinflammalory stimuli in MAECs to address their potential utility for investigation of disease mechanisms. LPS and cytokines induce leukocyte adhesion ligands and NO production (Marumo et al., 1993; Gerritsen et al., 1995; Geiger et al., 1997). Because VCAM-1 is present in several cell types (Miyake et al., 1991; Rosen et al., 1992), is inducible, and is a feature of macrovascular disease (Elices et al., 1990; Cybulsky and Gimbrone, 1991), we chose to investigate its expression in MAECs. Our results clearly showed that MAECs basally express VCAM- 1, that LPS induced it, and that IFN-[gamma] potentiated the inauction (Fig. 5). In contrast to VCAM-1, LPS alone did not affect NO production or iNOS protein levels in MAECs. However, the combination of LPS and IFN-[gamma] was highly synergistic in inducing both (Figs. 6 and 7). The synergistic effect of LPS plus IFN-[gamma] on NO production is reminiscent of the response of rat aortic endothelial cells (Marumo et al., 1993; Geiger et al., 1997). As expected, NO production was sensitive to the NOS inhibitors L- NMMA and aminoguanidine (Fig. 8). These results demonstrate that the MAECs retained typical responses to proinflammatory stimulation in culture.

In conclusion, the nonenzymatic method of explant culture, expansion, and selection for di-I-acetylated-LDL uptake allowed the reproducible isolation of pure cultures of aortic endothelial cells from few mice. The method is simple and convenient. The MAECs exhibited strong responses to the proinflammatory agents LPS and IFN- [gamma]. Thus, endothelial function and macrovascular disease mechanisms can be studied with these cells, taking advantage of useful murine models.

ACKNOWLEDGMENTS

We appreciate the assistance of Rustislav Likhotvorik and Andrew Oberyszyn (Ohio State University, Dorothy M. Davis Heart and Lung Research Institute) in conducting this work. This work was supported by an award from the American Heart Association, Ohio Valley Affiliate, 0151021B, and by National Institutes of Health Grants HL68054 and P30 CA16085.

In Vitro Cell. Dev. Biol.-Animal 39:43-50, January and February 2003

(C) 2003 Society for In Vitro Biology

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HONG HUANG, JANE MCINTOSH, AND DALE G. HOYT1

Division of Pharmacology, The Ohio State University College of Pharmacy, Columbus, Ohio 43210

(Received 20 March 2003; accepted 10 April 2003)

1 To whom correspondence should be addressed at Division of Pharmacology, The Ohio State University College of Pharmacy, 500 West Twelfth Avenue, Columbus, Ohio 43210. E-mail: hoyl.27@osu.edu

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