Journal of Histochemistry and Cytochemistry, Vol. 47, 159-168, February 1999, Copyright © 1999, The Histochemical Society, Inc.


ARTICLE

Caveolae and Vesiculo–Vacuolar Organelles in Bovine Capillary Endothelial Cells Cultured with VPF/VEGF on Floating Matrigel–collagen Gels

Eliza Vasilea, Qu-Honga, Harold F. Dvoraka, and Ann M. Dvoraka
a Departments of Pathology, Beth Israel Deaconess Medical Center and Harvard Medical School, Boston, Massachusetts

Correspondence to: Ann M. Dvorak, Dept. of Pathology, Beth Israel Deaconess Medical Center, 330 Brookline Ave., Boston, MA 02215.


  Summary
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In situ vascular endothelium is characterized by many cytoplasmic vesicles (caveolae) and vacuoles. In venules these are organized into prominent clusters called vesiculo–vacuolar organelles or VVOs. VVOs provide an important pathway for plasma protein extravasation in response to vasoactive mediators. In contrast, cultured endothelial cells isolated from many sources lack VVOs and generally have few caveolae. Our goal was to preserve VVOs in cultured endothelium. Bovine adrenal microvascular endothelial cells (BCEs) cultured on floating Matrigel–collagen Type I gels with vascular permeability factor/vascular endothelial growth factor (VPF/VEGF) exhibited typical VVOs by electron microscopy. Both in vivo and in culture VVOs were caveolin-positive by immunoelectron microscopy. On the basis of caveolin immunostaining, VVOs could also be detected by light (confocal) microscopy. When BCEs were cultured without VPF/VEGF, caveolin staining was finely punctate and electron microscopy confirmed the near absence of VVOs. BCE VVOs were sensitive to N-ethylmaleimide. Other types of endothelium cultured on Matrigel–collagen gels with or without VPF/VEGF exhibited few caveolae and no VVOs. Therefore, preservation of VVOs in cultured endothelium required a specific combination of endothelial cells (BCEs), surface matrix (Matrigel–collagen), and growth factor (VPF/VEGF). These endothelial cells should be useful for in vitro studies of trans-endothelial transport. (J Histochem Cytochem 47:159–167, 1999)

Key Words: caveolae, transcytosis, vesiculo–vacuolar organelle (VVO), vascular permeability factor/vascular endothelial growth factor (VPF/VEGF), caveolin, tissue culture, endothelial cells, Flk-1, ultrastructural immunocytochemistry, N-ethylmaleimide


  Introduction
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Introduction
Materials and Methods
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The vascular endothelium that lines most large and small blood vessels is characterized by many smooth cytoplasmic vesicles and vacuoles. Capillary endothelium is rich in homogeneously sized (60–80 nm) membrane-bound vesicles that have been termed plasmalemmal vesicles or caveolae (Palade and Bruns 1968 ). Most capillary caveolae are individual structures that invaginate from either the luminal or the abluminal plasma membrane. Sometimes, however, caveolae form aggregates of two or three interconnected vesicles and, less commonly, appear individually as free structures in the endothelial cell cytoplasm without connection to the plasma membrane. Capillary caveolae lack a visible cytoplasmic coat by electron microscopy, but a 21-kD transmembrane protein, caveolin-1 (identical to VIP-22) is associated with their cytoplasmic face (Rothberg et al. 1992 ).

Caveolae are believed to be responsible for transcytosis, the process by which plasma proteins are transported across capillary endothelium (Palade et al. 1979 ; Vasile et al. 1983 , Vasile et al. 1989 ; Vasile and Simionescu 1985 ; Ghitescu et al. 1986 ; Milici et al. 1987 ; Palade 1988 ; Schnitzer et al. 1995b ). Transcytosis is an active process in which caveolae that open to the vascular lumen take on plasma proteins as cargo. Such caveolae then separate from the luminal plasma membrane, pass across the endothelial cytoplasm, and fuse with the abluminal plasma membrane, where they discharge their contents into the underlying basal lamina. Transcytosis is strongly inhibited by N-ethylmaleimide (NEM), which alkylates NEM-sensitive factor (NSF), an ATPase that plays a key role in the fusion events of exocytosis and endocytosis (Predescu et al. 1994 ; Schnitzer et al. 1995a ). By analogy, transcytosis is believed to make use of mechanisms and proteins similar to those involved in the intracellular movements of other cytoplasmic vesicles, e.g., ER/Golgi transport vesicles and synaptic vesicles (Schnitzer et al. 1995b ; Predescu et al. 1997 ).

Venular endothelial cells in the skin, subcutis, skeletal muscle, and peritoneal lining tissues are taller than their counterparts in capillaries and contain in their cytoplasm prominent bunches of grape-like aggregates of interconnecting vesicles and vacuoles, termed vesiculo–vacuolar organelles or VVOs (Kohn et al. 1992 ; Dvorak et al. 1996 ; Feng et al. 1996 ). VVOs span venular endothelium from lumen to ablumen and are commonly composed of more than 100 individual vesicles and vacuoles. VVOs provide an important pathway for extravasation of circulating macromolecules in response to vasoactive agents such as vascular permeability factor/vascular endothelial growth factor (VPF/VEGF), histamine, or serotonin. VVOs are also prominent in the hyperpermeable endothelium of some tumor microvessels where they provide a pathway for plasma extravasation (Kohn et al. 1992 ; Feng et al. in press ).

In contrast to the microvascular endothelium in vivo, cultured endothelial cells derived from many different sources exhibit relatively few caveolae and generally lack VVOs. The reasons for this are not clear but may reflect a failure of complete endothelial cell differentiation in tissue culture. The paucity of vesicles and VVOs is an issue of some importance for studies of solute transport carried out in vitro on cultured vascular endothelium. If, as is generally accepted, macromolecules cross microvascular endothelium in vivo by way of caveolae or VVOs, then the absence of these structures would be expected to impair the transcytosis of such molecules across endothelial cell monolayers in culture. Indeed, transport of albumin and other solutes across endothelial cell monolayers is poorly responsive to vasoactive mediators such as histamine (reviewed in Curry 1992 ).

Because of our interest in macromolecular transport across endothelium in response to vasoactive agents, we sought to develop conditions for culturing vascular endothelium in which caveolae and VVOs were preserved. We here report progress toward this goal. Using floating gel matrices comprised of Matrigel and Type I collagen, along with appropriate concentrations of VPF/VEGF, we have been able to culture bovine capillary endothelial cells (BCEs) that contain many caveolae and prominent clusters of vesicles and vacuoles similar to the VVOs found in venular endothelium in vivo. Moreover, we have shown that the VVOs of cultured endothelium are sensitive to treatment with NEM and that they express caveolin-1 as do the VVOs found in venule endothelium.


  Materials and Methods
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Materials and Methods
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Reagents
Dulbecco's minimal essential medium (DMEM), fetal calf serum (FCS), and calf serum (CS) were purchased from JRH Laboratories (Lenexa, KS), endothelial growth medium (EGM) from Clonetics (San Diego, CA), recombinant human vascular endothelial growth factor (rVPF/VEGF) and basic fibroblast growth factor (bFGF) from R&D Systems (Minneapolis, MN), Matrigel from Becton Dickinson (Bedford, MA), and collagen type I from Collagen Biomaterials (Palo Alto, CA). Affinity-purified rabbit anti-caveolin polyclonal antibody was obtained from Transduction Laboratories (Lexington, KY) and rabbit anti-KDR antibody was the generous gift of Dr. D. Senger. Fluorescein isothyocyanate (FITC), Texas Red- and horseradish peroxidase (HRP)-conjugated secondary antibodies were obtained from Cappel Organon Teknika (Durham, NC), peroxidase-conjugated F(ab')2 antibodies from Amersham Life Sciences (Arlington Heights, IL), SuperSignal Chemiluminescent System from Pierce (Richmond, IL), and Vectashield from Vector Laboratories (Burlingame, CA). Electron microscopy reagents were purchased from Ted Pella (Reading, CA) and all other reagents from Sigma (St Louis, MO).

Endothelial Cells
Bovine capillary endothelial cells (BCEs) were a gift from Dr. Judah Folkman (Children Hospital Medical Center, Boston, MA). Human dermal microvascular endothelial cells (HDMVECs) were isolated and purified from human foreskin (Detmar et al. 1990 ). Human microdermal endothelial cells-1 (HMEC-1) were the gift of Dr. Thomas Lawley (Emory University, Atlanta, GA), and human umbilical endothelial cells (HUVECs) were purchased from Clonetics.

Tissue Culture
BCEs, passage 9–13, were plated either on gelatin-coated plastic dishes or on preformed Matrigel–collagen gels (Matrigel 10 mg/ml, collagen 1.5 mg/ml) and were grown in DMEM (low-glucose) supplemented with 10% CS, 5 U/ml penicillin, and 5 µg/ml streptomycin. BCEs were allowed to reach confluence on the preformed gels, after which the gels were detached from the plastic dish with a thin plastic spatula. The cells were then incubated for an additional 16 hr with or without 500 ng/ml rVPF/VEGF. Some of the VPF/VEGF-treated cells were additionally incubated with 5 mM NEM at 37C for 15 min immediately before fixation for immunofluorescence or electron microscopy. HMEC-1, HUVECs, and HDMVECs were grown on the same type of matrices as BCEs but in EGM supplemented with 10% FCS and 1 ng/ml hydrocortisone.

Immunofluorescence
For these studies, BCEs were grown as described above but on either preformed Matrigel–collagen gels cast on coverslips (400 µl/well) and then gently detached as above or on gelatin-coated coverslips in 12-well dishes. Cells were fixed with 4% formaldehyde in PBS for 1 hr at room temperature (RT) and then were permeabilized for 30 min with 0.1% Triton X-100 in PBS containing 1% BSA. Cells were then incubated with 2.5 µg/ml rabbit anti-caveolin antibody (primary antibody), washed in PBS, and incubated with 1:500 FITC-conjugated goat anti-rabbit IgG (secondary antibody). After further washing with PBS, the floating gels with attached cells were gently transferred to coverslips and mounted on microscope slides with Vectashield, a fluorescence anti-fading agent. Cells grown on gelatin-coated coverslips were prepared in the same manner. As negative controls, normal rabbit IgG at the same protein concentration was used instead of first antibody or the first antibody was omitted (Anderson et al. 1978 ). The cells were viewed with a Bio-Rad (Richmond, CA) MRC-1024 confocal microscope.

Electron Microscopy
BCEs grown on floating Matrigel–collagen gels in six-well dishes or on gelatin-coated plastic dishes were fixed with 2.5% glutaraldehyde and 2% formaldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, for 1 hr, postfixed in 1% OsO4 in 0.1 M sodium cacodylate buffer, pH 7.4, for 1 hr, stained en bloc with uranyl acetate, and embedded in Eponate. Thin Eponate sections were visualized with a Philips 300 electron microscope operated at 60 kV.

Immunoelectron Microscopy
BCEs grown on floating gels or on gelatin-coated plastic dishes were fixed with 4% formaldehyde in 0.05 M sodium phosphate buffer (PB), pH 7.2, for 1 hr and washed for 30 min in PB. Cells were then permeabilized in PB containing 0.05% Triton X-100 and 1% BSA for 60 min at RT. They were then incubated with first antibody (affinity-purified rabbit anti-caveolin) for 16 hr at 4C. The cells were washed in PB, incubated with HRP-conjugated secondary antibody for 60 min, and washed for 30 min in PB. Cells were then postfixed in 3% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, for 1 hr and washed in 0.05 M Tris-HCl buffer, pH 7.2. The peroxidase reaction was carried out by incubation in 0.2% 3,3'-diaminobenzidine (DAB) and 0.5% H2O2 in 0.05 M Tris-HCl buffer, pH 7.2, for 30 min at RT. Cells were then postfixed in reduced osmium (1% OsO4 and 1% potassium ferrocyanate in 0.1 M sodium cacodylate buffer, pH 7.4) for 1 hr and further processed for Eponate embedding as above. As negative controls, the first antibody was omitted or was replaced with normal rabbit IgG (Vasile et al. 1983 ). Guinea pig flank skin was processed for immunocytochemical detection of caveolin in situ in guinea pig skin venules by a similar methodology (Qu-Hong et al. 1995 ).

Immunoblotting
BCEs grown to confluence were incubated without VPF/VEGF or with VPF/VEGF for 1 or 16 hr. Cells were then washed in cold PBS containing 1 mM PMSF, detached from the matrix by incubation with PBS supplemented with 10 mM EDTA and protease inhibitors (10 µg/ml aprotinin, 10 µg/ml leupeptin, 10 µg/ml pepstatin, and 1 mM PMSF), and then centrifuged at 1000 x G for 10 min at 4C. Cell pellets were resuspended in the same solution and saved in aliquots. Aliquots for immunoblotting were centrifuged at 1000 x G at 4C. For detection of KDR (or VEGFR-2, one of the VPF/VEGF receptors and the human equivalent of murine flk-1), protein cell pellets were solubilized in a modified RIPA buffer (0.01 M Tris buffer, pH 8.0, 0.15 M NaCl, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS and protease inhibitors) for 10 min at 4C, briefly sonicated, centrifuged at 13,000 x g for 10 min to remove debris, and the supernatant mixed with Laemmli buffer and boiled for 5 min before electrophoresis.

For detection of caveolin, equivalent amounts of cell pellets (based on protein concentration) were dissolved and boiled in Laemmli sample buffer containing 2 M urea. (Laemmli 1970 ). Samples were subjected to 4–20% gradient SDS-PAGE, blotted onto 0.2-µm pore size PVDF (polyvinylidine difluoride) membranes (Bio-Rad), and blocked with 5% fat-free milk in TBS-T (0.01 M Tris buffer, pH 7.5, 0.15 M NaCl, 0.1% Tween-20) overnight at 4C. Immunoblots were incubated with 10 µg/ml rabbit affinity-purified antibody to KDR or to caveolin, then with peroxidase-conjugated secondary antibodies diluted 1:5000. Signal was detected and visualized using the SuperSignal Chemiluminescent System assay with HYPER-film ECL (Amersham Life Sciences).


  Results
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Materials and Methods
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Induction of Caveolae and VVOs in Cultured Endothelium
In contrast to vascular endothelium in situ, cultured endothelial cells typically exhibit few caveolae and no VVOs. We have previously failed in repeated attempts to preserve these structures in vitro. For example, cultured HDMVECs (passage 5–10), HMEC-1, or HUVECs variously plated on gelatin, on collagen gels, or on fibrin gels, either at low or high cell density in DMEM or EGM supplemented with 20% fetal calf serum with or without VPF/VEGF, exhibited few caveolae and no VVOs. The only exception was that modest numbers of caveolae were observed in HMEC-1 cells grown between layers of fibrin gel; however, no vesiclo–vacuolar aggregates resembling VVOs were detected.

We now report success in inducing VVOs in one type of endothelium (BCEs) cultured on floating gels composed of Matrigel and collagen I in the presence of VPF/VEGF. By light microscopy, BCEs grown to near confluence under these conditions exhibited a densely packed cobblestone appearance, similar to that of cultured large vessel endothelium (not shown). By electron microscopy, these cells expressed a normal complement of cytoplasmic organelles, including a well-developed Golgi area, free ribosomes, strands of rough endoplasmic reticulum, and well-preserved mitochondria (Figure 1). In addition, many vesicles and vacuoles were present in the cytoplasm (Figure 1B–G). In favorable sections, individual vesicles and vacuoles were observed to interconnect with each other and with the apical, lateral, and basal plasma membranes by means of stomata, many of which were closed by diaphragms. Sometimes interconnecting vesicles formed tubular structures (Figure 1C, asterisks). BCE sometimes extended processes into the underlying Matrigel–collagen I matrix, and these displayed many uniformly sized caveolae that opened to or were in close proximity to the plasma membrane (Figure 1D). In some areas, endothelial cells were greatly thinned such that a single vacuole spanned the cytoplasm from apex to base (Figure 1F, arrowhead). Less commonly, thinned endothelium developed typical fenestrae that were guarded by diaphragms (Figure 1G), similar to those found in the fenestrated microvascular endothelium of endocrine glands, and of kidney and other tissues.



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Figure 1. Electron micrographs of BCEs cultured on floating Matrigel–collagen I gels with VPF/VEGF. (A) Perinuclear area of a cultured BCE shows well-preserved ultrastructure with characteristic cytoplasmic organelles, including Golgi zone with flattened cisternal stacks, free ribosomes, and rough endoplasmic reticulum. bl, basal lamina. (B) Thinned portion of peripheral cytoplasm displays many cytoplasmic vesicles and vacuoles, some of which are obviously interconnected and many of which open to the apical or the basal plasma membrane. L, nominal lumen. (C) Large clusters of interconnected vesicles and vacuoles form characteristic vesiculo–vacuolar organelles (VVOs) which span the entire cell cytoplasm from apex (nominal lumen) to basal (abluminal) surface. Some of the interconnected vesicles are tubular (asterisks). (D) Cytoplasmic processes that extend into the Matrigel–collagen matrix, as shown here, have many peripheral vesicles (caveolae) that open to or are located close to the plasma membrane. Note many central parallel filaments. (E) Typical VVO in peripheral cytoplasm traverses endothelium from apical to basal surface. (F) An attenuated portion of endothelial cytoplasm is spanned from apex to base by a single vacuole (arrowhead). (G) Two overlapping endothelial cells. One (top) has many cytoplasmic vesicles and vacuoles; the other (lower) is attenuated, with few vesicles and two fenestrae (arrows). Bars = 0.1 µm.

Caveolin Immunostaining of VVOs
To obtain evidence for a possible relationship between caveolae and VVOs, we performed electron microscopic immunocytochemistry to determine whether VVOs expressed caveolin-1, a protein known to be associated with caveolae in capillary endothelium (Rothberg et al. 1992 ). The results obtained (Figure 2A–C) indicate that VVOs (as well as caveolae) in both cultured BCE and in normal guinea pig skin strongly express caveolin-1. In addition, focal patches of plasma membrane were also caveolin-positive (Figure 2B, arrowhead), as has also been described in other cell types (Smart et al. 1994 ). Presumably, these patches of plasma membrane are related to caveolae in some way, perhaps serving as caveolae precursors.



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Figure 2. Electron microscopic immunocytochemistry reveals caveolin-positive vesicles and vacuoles in BCEs grown on a floating Matrigel–collagen I gel with VPF/VEGF (A,B) and in situ in a venule of guinea pig skin (C). (A) Two overlapping endothelial cells, the uppermost of which displays many caveolae and clusters of vesicles and vacuoles (VVOs) (bracket) that are decorated with enzyme reaction product. (B) Caveolae (arrows), VVOs (curved arrows), and a short segment of plasma membrane (arrowhead) are strongly caveolin-positive. (C) VVO and caveolae of venular endothelium of normal guinea pig skin are strongly caveolin-positive. L, lumen. Bars = 0.2 µm.

We next employed the same anti-caveolin antibody to determine whether we could identify VVOs by confocal microscopy. BCEs grown on floating Matrigel–collagen gels in the presence of 500 ng/ml VPF/VEGF exhibited large, caveolin-positive cytoplasmic aggregates that were especially prominent at the cell periphery (Figure 3A–C). These aggregates were present in almost every cell (Figure 3A) and followed the same distribution as the VVOs we observed in these cells by electron microscopy (Figure 1B–G). In some cells, aggregates were confined to a few focal areas at the cell periphery (Figure 3B), whereas in others they were more numerous and were distributed more uniformly (Figure 3C). Treatment of BCEs with 5 nM NEM for 15 min at 37C immediately before fixation greatly reduced caveolin-positive cytoplasmic aggregates (Figure 3D). By electron microscopy, NEM-treated cells were not injured and exhibited caveolae but very few vesicle clusters (not shown). In addition, BCEs grown with VPF/VEGF on gelatin-coated surfaces (instead of on floating Matrigel–collagen I matrices) displayed a fine, punctate caveolin staining pattern but did not reveal caveolin-positive aggregates characteristic of VVOs.



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Figure 3. Confocal fluorescence images of caveolin immunostaining in BCEs grown on floating Matrigel–collagen I gel with (A–D) or without (E) VPF/VEGF. (A) Low-magnification overview of BCEs cultured with VPF/VEGF shows many cytoplasmic aggregates of caveolin-positive structures, the majority of which are localized to the cell periphery. (B,C) Higher-magnification views of BCE cells cultured with VPF/VEGF displaying prominent, caveolin-positive aggregates in the cytoplasmic periphery (B) or more uniformly distributed in the cytoplasm (C). (D) BCEs as in A–C but treated with NEM for 15 min before fixation. Note loss of most caveolin-positive aggregates, although one remains (lower). Caveolin staining appears reduced overall and is dispersed finely in the cytoplasm. (E) BCEs cultured on floating Matrigel–collagen I gel but without VPF/VEGF. Caveolin staining is for the most part finely punctate and evenly distributed through the cytoplasm, although a few aggregates are present. Bars = 5 µm.

When BCEs were cultured on floating Matrigel–collagen I gel matrices but without VPF/VEGF, caveolin staining was more diffuse and finely punctate, as has been described in other cells containing caveolae both in vivo and in vitro (Smart et al. 1994 ), although occasional larger aggregates were also present (Figure 3E). A diffuse caveolin staining pattern was also observed when BCEs were grown on gelatin-coated surfaces, but the intensity and extent of staining were considerably less (not shown). Immunoblotting revealed no significant change in caveolin or in KDR (VEGFR-2) protein content in BCEs cultured with or without VPF/VEGF (Figure 4).



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Figure 4. Immunoblots for detection of KDR and caveolin-1 in BCE cells grown without (Lane 1) or with (Lanes 2 and 3) VPF/VEGF for 1 and 16 hr. Cells were solubilized in RIPA buffer for KDR detection and in SDS lysis buffer for caveolin detection. Solubilized extracts were subjected to SDS-PAGE (20 µg protein/lane for caveolin, 50 µg protein/lane for KDR) and then to immunoblot detection with antibodies to KDR and caveolin-1. Levels of KDR and of caveolin-1 protein did not change detectably when cells were grown with or without VPF/VEGF.


  Discussion
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Materials and Methods
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Caveolae and VVOs are generally infrequent or absent when endothelial cells from a variety of sources are grown in tissue culture. The work presented here has defined a set of conditions for preserving caveolae and VVOs in cultured BCE cells. That specialized culture conditions should be necessary for preserving these structures, characteristic of differentiated vascular endothelium, is not surprising. Many previous authors have called attention to the importance of matrix for maintaining the differentiated state of both cultured epithelium (Emerman and Pitelka 1977 ; Streuli and Bissell 1991 ) and endothelial cells (Folkman and Haudenschild 1980 ; Davies et al. 1984 ; Furie et al. 1984 ; Madri et al. 1988 ; Beck and D'Amore 1996 ; Esser et al. 1998 ). Failure to differentiate completely might be attributable to many factors, including loss of cell polarity, failure to synthesize an appropriate basal lamina, unphysiological concentrations of growth factors, and lack of mechanical stimulation or pressure gradients as occur in vivo (Davies et al. 1984 ; Palade 1988 ; Ott et al. 1995 ).

Unfortunately, conditions that permitted the preservation of abundant caveolae and VVOs in cultured BCEs were not successful in preserving these structures when applied to other types of cultured endothelium. Our failure cannot be attributed to the absence of such structures in the starting cells. For example, in vivo dermal microvessels from which early passage HUVECs were derived are rich in both caveolae and VVOs (Kohn et al. 1992 ; Dvorak et al. 1996 ; Feng et al. 1996 ). The properties of BCE that allow them, but not the other types of endothelial cells we tested, to develop caveolae and VVOs in culture are not yet known. In light of our results, it appears reasonable to believe that the preservation and induction of caveolae and VVOs are highly dependent on the composition and perhaps other properties, such as elasticity, of the extracellular matrix, expression of VEGF receptors, presence of VEGF, and perhaps other as yet undefined cytokines. Whatever the factors involved, the ability to induce caveolae and VVOs in cultured BCEs is important because caveolae are believed to be essential for the transcytosis of macromolecules across capillary endothelium (Simionescu 1983 ; Vasile and Simionescu 1985 ; Palade 1988 ; Vasile et al. 1989 ), and VVOs have been shown to have an important role in macromolecular extravasation across venules and tumor microvessels in response to VPF/VEGF and other vasoactive mediators (Kohn et al. 1992 ; Dvorak et al. 1996 ; Feng et al. 1996 ; Feng et al. in press ). Therefore, previous studies of macromolecular extravasation across monolayer endothelial cell cultures may need to be reexamined because the cultured endothelial cells used very probably had few or no caveolae or VVOs (reviewed in Curry 1992 ).

A second new finding is that the VVOs of cultured BCEs, as well as those found in situ in the venular endothelium of normal skin, are caveolin-positive, as demonstrated by electron microscopic immunocytochemistry. These data add to the evidence supporting a relationship between VVOs and caveolae. Other evidence includes the finding that the smaller vesicles of VVOs are identical in morphology to caveolae and that VVO vacuoles have volumes that are not continuous but rather correspond to multiples of the volume of caveolae (Feng et al. in press ).

A third important finding is that, because of their caveolin-positivity and arrangement in aggregates, VVOs have been visualized by confocal microscopy. This should provide a useful screening procedure for tentatively identifying VVOs in cultured endothelium without the need for electron microscopy.

Finally, we have demonstrated that VVOs are reduced by brief exposure to NEM, as demonstrated both by electron microscopy and by loss of caveolin-positive aggregates by confocal microscopy. NEM is an alkylating agent which, among other functions, interferes with caveolae-mediated transcytosis (Predescu et al. 1994 ; Schnitzer et al. 1995a ). It is tempting to speculate from this finding that NEM disrupts transcytosis in vivo by disrupting VVOs.

Recently, Esser et al. 1998 reported that BCEs developed many fenestrae when they were co-cultured with epithelial cells that had been stably transfected to express VPF/VEGF or on epithelial cell-derived matrices in the presence of added VPF/VEGF. In addition, these BCE also developed some "large clusters of fused vesicles" that were believed to be comparable to the VVOs that we have described in vivo (Kohn et al. 1992 ; Dvorak et al. 1996 ; Feng et al. 1996 ) and that we have reported here. Similar to our results, Esser et al. 1998 found that generation of "clustered vesicles" (VVOs) required a particular endothelial cell (BCE), an appropriate basal lamina-type matrix (in their case generated by cultured corneal endothelial cells), and VPF/VEGF (either provided by co-culture with VPF/VEGF-transfected epithelium or by added recombinant VPF/VEGF). However, despite these similarities to our results, the vesicle clusters described by Esser et al. were caveolin-negative with an immunogold postembedding protocol. In contrast, the VVOs we studied in vivo and in vitro were, like caveolae, strongly caveolin-positive, as determined with a pre-embedding immunoperoxidase method. Another difference in results is that the cells described by Esser et al. were highly fenestrated, whereas fenestrae, although occasionally present, were not common in our cultured BCEs.

In summary, we have defined a relatively simple culture procedure for growing endothelial cells in a manner that preserves VVOs and caveolae, properties characteristic of differentiated endothelium in vivo. Endothelial cells cultured in this manner should be useful in future studies of transendothelial cell transport of plasma proteins and other solutes.


  Acknowledgments

Supported by US Public Health Service National Institutes of Health grants CA-50453 and AI-33372.

Received for publication August 7, 1998; accepted September 30, 1998.


  Literature Cited
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Materials and Methods
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