Nonuniformity of Endothelial Constitutive Nitric Oxide Synthase Distribution in Cardiac Endothelium

From the Department of Physiology, University of Antwerp (Belgium).

Correspondence to S.U. Sys, Department of Physiology, University of Antwerp (RUCA), Groenenborgerlaan 171, B-2020 Antwerpen, Belgium. E-mail STSYS{at}RUCA.UA.AC.BE

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Abstract—Endocardial endothelium and endothelium of coronary vessels produce NO. Histochemical methods have suggested that coronary arterial endothelial cells contain more endothelial constitutive NO synthase (ecNOS) than does coronary venous endothelium. We have further investigated the distribution of ecNOS in cardiac endothelium using immunofluorescence and en face confocal microscopy of rat heart. In endocardial endothelium, confocal microscopy revealed distinct ecNOS labeling of peripheral cell borders, cytoplasmic labeling, and labeling of the Golgi complexes. Labeling of the cell borders and of the Golgi complexes was confirmed by double staining for ecNOS and for platelet and endothelial cell adhesion molecule or Golgi 58k protein, respectively. Cytoplasmic labeling was strongest in coronary arterial endothelium. The size of the ecNOS-labeled Golgi complexes decreased from coronary arterial endothelial cells (8.63±0.39 µm², mean±SE of 5 rats) to endocardial endothelium (7.07±0.61 µm²) and to coronary venous endothelium (3.65±0.20 µm²). In addition, pixel intensity of ecNOS labeling was higher in arterial endothelial cells than in venous endothelial cells. Endothelium of myocardial capillaries also contained small ecNOS-labeled Golgi complexes. No correlation was observed between endothelial cell surface area and Golgi complex size. Caveolin-1 labeling was strongest in capillaries and did not coincide completely with ecNOS labeling in endocardial and venous endothelium. These results suggest that endocardial and coronary arterial endothelium in the rat have a higher synthetic activity and might express more ecNOS than is expressed by cardiac venous and capillary endothelium. The observed heterogeneity in ecNOS distribution might be related to the specific mechanochemical environment and function of each endothelial compartment.

Key Words: constitutive nitric oxide synthase • endocardial endothelium • coronary endothelium • Golgi complex • caveolin

	Introduction

Top Abstract Introduction Materials and Methods Results Discussion References

 
Nitric oxide is a widespread biological mediator that is implicated in various physiological and pathophysiological processes. In the heart, NO modulates not only coronary flow but also myocardial performance. This latter feature has been shown in several species,^{1 2 3 4 5} including humans.⁶ Whereas higher concentrations of NO produce a negative inotropic effect, at low concentrations it can induce a distinct positive inotropic effect.⁵ In physiological conditions, NO is synthesized from L-arginine by constitutive NOS. The two known isoforms of constitutive NOS have been observed in the heart.

The neuronal isoform (nNOS or NOS type I) is present in a subpopulation of intracardiac ganglia and nerve fibers throughout atrial tissue⁷ and in some perivascular nerve fibers of ventricular myocardium.⁸ When the NADPH-diaphorase method is used to localize constitutive NOS in the ventricle of pig heart, most staining has been found in coronary vascular endothelium and in the endocardium.⁸ Immunohistochemical methods confirmed the presence of the endothelial constitutive NOS isoform (ecNOS or NOS type III) in cardiac endothelium.^{9 10} Compared with ecNOS labeling in the endothelium of coronary vessels, a modest ecNOS labeling has also been observed in cardiac myocytes,⁴ where it appears to be associated with caveolin-3,¹¹ a muscle-specific isoform of a coat protein of caveolae.

In ECs, ecNOS is associated primarily with the particulate fraction,^{12 13 14} in particular with the Golgi complex, and with domains of the plasma membrane, the caveolae.^{12 13 15 16 17} The Golgi complex (also called Golgi apparatus) consists of one or more stacks of cisternae surrounded by vesicles.¹⁸ The Golgi complex is the site of biosynthesis of glycolipids and of sugar moieties of glycoproteins.¹⁸ The processed proteins, lipids, and polysaccharides are either directly secreted or sorted in the Golgi complex and transported to other cell organelles, cell membranes, secretory granules, and lysosomes. Caveolae are specialized invaginations of the plasma membrane enriched with caveolin-1 (the nonmuscle isoform of a coat protein of caveolae), Ca²⁺-ATPase, G proteins, and inositol trisphosphate receptors.^{19 20 21} Recently, enzymatic ecNOS activity was demonstrated in caveolar membranes.¹⁶ ecNOS colocalized with caveolin-1 in cultured microvascular cells,²² and antibodies to caveolin-1 immunoprecipitated ecNOS from the endothelium of myocardial capillaries.¹¹ In addition to their role in endocytosis, the caveolin-rich membrane domains are thus also engaged in signal transduction.

Although ecNOS is a constitutive enzyme, its expression in vascular endothelium can be modulated by shear stress,²³ oxygen,²⁴ transforming growth factor-ß,²⁵ cytokines,²⁶ and various other factors (for a review see References 27 and 28^{27 28} ). Regional differences in these modulating conditions might explain the wide variations in expression of ecNOS observed in endothelium of various cardiovascular segments. Heterogeneity of immunohistochemical ecNOS staining in endothelium has been reported in lung and renal vessels.^{29 30} In rat heart, NADPH-diaphorase staining was more intense in coronary arterial ECs than in venous ECs.⁸ It is not known whether ecNOS labeling of endocardial ECs and other cardiac ECs displays differences in its intensity or intracellular distribution.

In the present study, we have demonstrated, through whole-mount immunostaining and en face confocal microscopy, that ecNOS was present in all cardiac ECs. Staining was more intense in arterial and endocardial endothelium than in the endothelium of coronary veins and myocardial capillaries. This difference in staining was associated with heterogeneity in the size of the Golgi complexes.

	Materials and Methods

Top Abstract Introduction Materials and Methods Results Discussion References

 
For en face confocal laser scanning microscopy, rats were perfused with 2% paraformaldehyde in a HEPES buffer (mmol/L: NaCl 136.9, KCl 2.7, MgCl₂ · 6H₂O 0.5, KH₂PO₄ 1.50, NaH₂PO₄ · H₂O 8.10, glucose 5.0, and HEPES 20, pH 7.2; total fixation time of 20 minutes: 5 minutes at 37°C and 15 minutes at room temperature). Strips of endocardium (right ventricle: septum and tendon end of papillary muscles, septum, outflow tract, and pulmonary valve; left ventricle: papillary muscle, septum, and outflow tract), myocardium (right and left ventricular free wall), and coronary vessels were isolated and permeabilized with 0.2% Triton X-100 in PBS. Permeabilization and all following staining steps were performed in Eppendorf vessels. After rinsing with PBS-BSA (0.2% BSA in PBS) and immersion for 30 minutes in blocking solution (0.2% BSA and 5% goat serum in PBS or 12.5 µg purified goat IgG/mL), the tissue samples were incubated for 1 to 2 hours in a PBS-BSA solution containing the primary antibody. In single whole-mount staining experiments, primary monoclonal and polyclonal antibodies were used to detect the expression of ecNOS, nNOS, or caveolin-1 (a marker of noncoated vesicles in nonmuscle cells). Double staining was used to investigate the colocalization of ecNOS and PECAM, Golgi 58k protein (a Golgi marker³¹ ), or RECA-1. For this purpose, tissue strips were immersed in a mixture of polyclonal ecNOS and PECAM antibodies (or Golgi 58k protein or RECA antibodies). The Table lists the origin of the primary antibodies and the working dilution. For single immunofluorescence labeling, goat anti-mouse or goat anti-rabbit antibodies coupled to DTAF (dilution, 1/100, Jackson Laboratories) were used as secondary antibodies. For double staining, specimens were incubated with a mixture of goat anti-mouse antibody coupled to DTAF and goat anti-rabbit antibody coupled to indocarbocyanine (dilution, 1:100; Jackson Laboratories). Single and double staining were also performed on cryostat sections of rat heart. Perfusion-fixed rat hearts were immersed for 2 hours in a 40% sucrose solution, mounted in OCT (Tissue Tek), and frozen in liquid nitrogen.

En face preparations and cryostat sections were mounted in Slowfade Light (Molecular Probes). Coverslips were sealed with nail polish. Both preparations were observed with a Polyvar 2 epifluorescence microscope and with a Bio-Rad 600 confocal laser scanning microscope. For confocal microscopy, the BHS filter block was used for single stainings. The A1 and A2 filter blocks were used for observing double stainings. In negative controls (for which only the secondary antibodies were used), no staining was observed in whole mounts or cryostat sections. Image files were stored on optical disks and further viewed with Confocal Assistant (CAS), a free-ware program made by Todd Clark Brelje. Image processing and analysis of the Bio-Rad images was performed with Fenestra, a Windows-based program. Morphometric data were exported to Excel and SPSS for statistical analysis.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
Endocardial Endothelium
En face confocal microscopy of endocardial strips from different zones, including the cardiac valves, of left and right ventricle demonstrated that all endocardial ECs expressed ecNOS. The three primary ecNOS antibodies used in the present study yielded similar results. Staining with the polyclonal antibody from Transduction Laboratories had a more granular appearance. Immunofluorescence staining of endocardial ECs revealed cytoplasmic labeling that outlined the unstained nuclei. In each endocardial EC, a brightly stained spot corresponding to the Golgi complex was located near the nucleus (Fig 1A and 1B). Occasionally, one cell contained two Golgi complexes. Usually, a Golgi complex consisted of an area, with a ringlike or more complex shape, of intense labeling and an unstained or lightly stained central dot. Double staining with ecNOS and Golgi 58k protein, a Golgi marker, confirmed that ecNOS had indeed labeled the Golgi complex. The periphery of the endocardial EC was outlined by an ecNOS-stained bandlike structure coinciding with PECAM labeling in double-stained preparations (Fig 2A and 2B). PECAM is known to label the whole depth of endothelial intercellular clefts. In the heart, all PECAM-labeled ECs expressed ecNOS. The width of the PECAM bands (hence, of the intercellular clefts) was larger in endocardial ECs than in arterial and venous ECs, thus confirming previous ultrastructural observations.³² In several endocardial ECs, PECAM labeling was also observed in Golgi complexes.

View larger version (137K): [in this window] [in a new window]   Figure 1. En face confocal microscopy showing distribution of ecNOS in cardiac endothelium. Compared with those for endocardial ECs and arterial ECs, contrast settings and size of the confocal pinhole were increased to enhance images of ecNOS labeling in the endothelium of myocardial capillaries and of coronary veins. EE indicates endocardial endothelium; AE, arterial endothelium; and VE, venous endothelium. A and B, In endocardial ECs, ecNOS labeling was distinct in Golgi complexes (arrows) and along the cell periphery (arrowheads). Cytoplasmic labeling outlined the nuclei (Nu). Bars=20 µm (A) and 10 µm (B). C and D, Golgi complexes were more intensely labeled and more elongated in the endothelium of coronary arteries than in endocardial ECs. Arterial ECs were not outlined by peripheral ecNOS staining. Only some parts of the cell periphery were distinctly stained (arrowheads). Cytoplasmic labeling outlined endothelial nuclei (Nu). Bars=20 µm (C) and 10 µm (D). E, ecNOS labeling and the shape and size of Golgi complexes in the endothelium of a coronary arteriole were similar to those in AE. Bar=10 µm. F, In the endothelium of myocardial capillaries, Golgi complexes were small, and cytoplasmic ecNOS labeling was weak. Labeling of intercellular borders was not observed. Bar=10 µm. G and H, In coronary veins, Golgi complexes (arrows) were small and sometimes below the nucleus (arrows in panel H). Cytoplasmic ecNOS staining was rather weak, except along the narrow endings of one or both sides of ECs (arrowheads in panel G). A thin band of ecNOS labeling outlined many but not all venous ECs. Bars=20 µm (G) and 10 µm (H).

View larger version (87K): [in this window] [in a new window]   Figure 2. En face confocal microscopy and double immunolabeling of cardiac endothelium. EE indicates endocardial endothelium; AE, arterial endothelium. A and B, PECAM-1 and ecNOS labeling. In endocardial ECs, the peripheral PECAM- labeled band coincided with the peripheral ecNOS. PECAM labeling of Golgi complexes was usually observed in EE. Bar=10 µm. C and D, Golgi 58k protein and ecNOS labeling. Thin optical section through endothelium of a coronary artery shows coincidence of the Golgi marker with ecNOS staining. Bar=10 µm. E and F, RECA-1 and ecNOS labeling. Thick optical section through ventricular myocardial tissue is shown. All RECA-labeled microvascular ECs were also labeled for ecNOS. Bar=20 µm.

Immunostaining for caveolin-1, with monoclonal or polyclonal antibodies, showed that peripheral borders of endocardial ECs were nearly completely devoid of caveolin labeling (Fig 3A, 3B, and 3C). The pattern of these dark unstained peripheral areas was similar to PECAM-1 labeling. Many endocardial ECs also contained juxtanuclear caveolin labeling. Subjacent cardiomyocytes were not labeled (Fig 3C). The juxtanuclear labeling was more evident after staining with the monoclonal anti–caveolin-1 antibody.

View larger version (139K): [in this window] [in a new window]   Figure 3. Confocal microscopy showing distribution of caveolin-1 in cardiac endothelium. EE indicates endocardial endothelium; AE, arterial endothelium; and VE, venous endothelium. A and B, En face optical sections through endocardial ECs reveal the absence of labeling along the cell periphery (arrowheads). Nuclei (Nu) are distinctly outlined by juxtanuclear labeling and by labeling of the cell membrane. C, Optical section through a small trabeculum is shown. Cardiomyocytes in the central core of the trabeculum are unstained. Endocardial cells contain luminal and abluminal labeling. Labeling is weak or absent near the cell periphery (arrowheads). D, En face optical section through arterial ECs is shown. Thin unstained bands (arrowheads) outline the periphery of the arterial ECs. Cytoplasmic and membranous labeling show a pattern distinct from that in endocardial ECs. E, Caveolin-labeling of endothelium of myocardial capillaries is more intense than in other cardiac ECs. It is not clear whether the dark lines within the capillaries represent the peripheral borders or other structures. F, En face section through venous ECs is shown. The elongated cell shape of the venous ECs is evident by the absence of caveolin-staining along the cell periphery (arrowheads). Bars=10 µm.

Although nNOS labeling was not detected in endocardial ECs, it was prominent in neurons and nerve fibers of ganglia in the right atrium near the superior vena cava (not shown).

PECAM and ecNOS labeling revealed considerable differences in endocardial EC size between various areas of the right and left ventricular endocardial endothelium. In some hearts, larger endocardial ECs, which were usually observed in the right ventricular outflow tract and septum, possessed a larger Golgi complex than the small endocardial ECs, which can be found on the tendon end of right ventricular papillary muscles or on left ventricular papillary muscles. In other hearts, however, the size of the Golgi complexes and the degree of cytoplasmic staining did not differ between various areas within and between the right and left ventricles.

Endothelium of Coronary Arteries
Optical sections through coronary vascular endothelium (Fig 1C and 1D) revealed a considerably different pattern of ecNOS staining than did sections through endocardial ECs. In low-power images, there was an absence of the staining of peripheral cell borders in arterial endothelium. Double staining with ecNOS and Golgi 58k protein confirmed that the most brightly labeled structures in arterial ECs (Fig 2C and 2D) coincided with Golgi complexes. The Golgi complexes were usually located alongside the nucleus. As in endocardial ECs, a dark unstained area was present in the Golgi complexes of arterial ECs. Golgi complexes had a more elongated and frequently more complex shape in arterial endothelium than in endocardial ECs. In several arterial ECs, the Golgi complex consisted of nearly disconnected granule-like spots. In the endothelium of arterioles (Fig 1E), Golgi complexes had a shape and size similar to those in the endothelium of coronary arteries. On the other hand, ecNOS labeling of endothelium of the thoracic aorta had a pattern similar to that in endocardial ECs. Golgi complexes were somewhat smaller in aortic ECs (6.45±0.25 µm², mean±SE, n=53) than in coronary arterial ECs (8.40±0.23 µm², n=77; both mean±SE values from aorta and coronary artery were from the same rat).

Double-labeled preparations of coronary arteries and aorta demonstrated that each of the PECAM-labeled cells was ecNOS positive. In addition, double labelings of arterial ECs showed that peripheral PECAM staining usually did not coincide with ecNOS staining (Fig 4C). In nearly all cells, the thin PECAM bands retained a green color in merged images, thereby outlining the red-colored ecNOS staining of cytoplasm and Golgi complexes. In several arterial ECs, PECAM-labeling surrounded linearly arranged ecNOS-positive structures, which were difficult to recognize as Golgi complexes in single ecNOS stainings. Only occasionally, parts of the periphery of arterial ECs were ecNOS positive. In contrast, like endocardial ECs, aortic ECs also showed ecNOS-positive peripheral borders.

View larger version (127K): [in this window] [in a new window]   Figure 4. A and B, Double immunolabeling of RECA and ecNOS of a cryostat section through the left ventricular wall. All microvessels labeled by RECA were ecNOS positive. C, En face optical section through arterial EC (arterial endothelium [AE]) after double labeling for PECAM-1 (green color) and ecNOS (red color). The green color of the peripheral bands shows the absence of colocalization with ecNOS. Areas where both stains colocalize should have a yellow color. Bar=10 µm. D through G, Confocal images from ecNOS-stained endocardial ECs (D and E), coronary arterial ECs (F), and venous endothelium (VE) (G) obtained by the same contrast and pinhole settings of the confocal microscope. In panel D, pixel-gray values were color-coded showing distinct Golgi complexes and peripheral bands. Other areas of the endocardial ECs (endocardial endothelium [EE])were weakly labeled. Note the differences in intensity of labeling between various endocardial ECs, which mainly resulted from different levels of focusing. In panels E and F, the pixel-gray values were color- and height-coded to appreciate better the differences in labeling intensity of the Golgi complexes in cardiac endothelium. High and yellow to white areas indicate zones of intense labeling. The peaks in arterial ECs (F), representing Golgi complexes, were larger and had pixel values higher than the peaks in endocardial ECs (E) and even higher than the peaks in venous ECs (G). In the cytoplasm between Golgi complexes, more green and light-blue pixel values were present in arterial endothelium than in endocardial ECs. Cytoplasmic labeling was weak in VE.

Caveolin-1 labeling of arterial ECs consisted of a patchy distribution of intense or weakly stained and unstained cellular areas (Fig 3D). Nuclei were more difficult to observe in arterial ECs than in endocardial ECs. The peripheral borders of arterial ECs were caveolin negative, but the unstained bands were much thinner than in endocardial ECs and sometimes difficult to discern by their irregular appearance and by the presence of unstained cellular zones. nNOS labeling was not detected in the endothelium of coronary arteries.

Endothelium of Myocardial Capillaries
Optical sections through myocardial tissue strips showed rather weak cytoplasmic ecNOS labeling of capillary endothelium. The ecNOS labeling outlined the nuclei (Fig 1F). The labeled Golgi complexes occupied a small area and usually had a circular or oval shape. Peripheral borders of the capillary ECs were not discernible. Double-stained preparations showed complete overlap of ecNOS labeling and RECA labeling of the ECs in myocardial microvessels (Fig 2E and 2F). RECA is an antibody that labels endothelium of the entire vasculature in all organs and tissues in the rat, including the sinusoidal endothelium and the high endothelium in lymphoid vessels.³³ Cryostat sections of myocardial tissue also showed complete overlap of ecNOS and RECA labeling of ECs in myocardial microvessels (Fig 4A and 4B). No differences in the pattern of ecNOS labeling of microvascular endothelium were observed between right and left ventricular myocardium. No specific staining was detected in cardiac myocytes. The degree of staining in cardiac myocytes after ecNOS labeling was similar to that in negative controls, where primary antibodies were omitted. In myocardial tissue strips, the polyclonal ecNOS antibody from Transduction Laboratories produced a higher aspecific background in cardiac myocytes and interstitial tissue than the other primary antibodies.

After PECAM-1 staining, capillaries were rather weakly labeled. In some capillaries, thin bands could be observed, suggesting the existence of thin peripheral borders. In contrast to PECAM-1 staining, immunostaining for caveolin-1 produced very intense labeling of myocardial capillaries (Fig 3E). Endothelium of myocardial capillaries was much more strongly stained for caveolin-1 than was endocardial or arterial endothelium. Adjacent cardiomyocytes were unstained.

Endothelium of Coronary Veins
Venous ECs were less intensely stained with ecNOS than were endocardial ECs and ECs of coronary arteries. A substantial increase of contrast and gain of the confocal microscope was necessary to visualize ecNOS staining in coronary veins. ecNOS-stained peripheral borders outlined venous ECs, displaying their typical elongated shape (Fig 1G). The cell borders were thin and weakly labeled; frequently, only one or both extremities of the cells were stained as intensely as the Golgi complexes (Fig 1H). Golgi complexes appeared small and had an unstained center. Golgi complexes were more frequently located underneath the nucleus in venous ECs than in endocardial ECs and arterial ECs. Caveolin-1 staining yielded similar images in venous ECs and arterial ECs. The unstained peripheral bands were more distinct in venous ECs than in arterial ECs (Fig 3F).

Image and Statistical Analysis
The intensity of cytoplasmic ecNOS staining in cardiac ECs was compared by using three-dimensional color-coded plots of pixel-gray values from images taken with the same contrast and gain settings and width of the confocal aperture on the Bio-Rad MRC-600 (Fig 4E through 4G). Golgi complexes in arterial ECs were not only larger but also more intensely labeled than Golgi complexes in endocardial ECs and especially in venous ECs. Between the Golgi complexes, more green-coded and light-blue–coded pixels were present in arterial ECs than in endocardial ECs. Venous ECs contained the least cytoplasmic labeling around the Golgi complexes.

The area of the Golgi complexes, detected by ecNOS staining, was nonuniformly distributed in cardiac ECs. The distributions of the area of Golgi complexes in endocardial (right ventricle and septum), arterial, and venous cells (n=5 rats, 701 measured areas) were significantly right-skewed. Logarithmic transformation was used to normalize these distributions; the variances were stabilized by the same procedure. Coefficients of variation for the endothelial Golgi complex area, derived from the within-rat variation in a one-way ANOVA on the transformed data, were similar (Levene test): 25.8%, 26.8%, and 26.8% for endocardial, arterial, and venous ECs, respectively. The means of backtransformed Golgi complex area were significantly different among endocardial, arterial, and venous ECs (Fig 5A). Measurements of Golgi complexes in the endothelium of myocardial capillaries yielded similar or smaller values than found in venous ECs.

View larger version (37K): [in this window] [in a new window]   Figure 5. Graphs showing data (mean±SE) obtained by morphometric image analysis of confocal images from endocardial ECs, arterial endothelium (AE), and venous endothelium (VE). EE indicates endocardial endothelium. A, The means of the area of Golgi complexes were significantly different in the three types of endothelium. The largest Golgi complexes were present in AE, and the smallest ones were in VE. *P<.01. B, The shape of Golgi complexes differed significantly in cardiac endothelium. The largest maximal cord was present in arterial ECs, and the smallest ones were in venous ECs. *P<.01. C, Plot shows the size of Golgi complexes and corresponding EC area (mean±SE) from coronary VE (open symbols), EE (closed symbols), and AE (shaded symbols) from four different hearts (square, rat 1; triangle, rat 2; inverted triangle, rat 3; and diamond, rat 4). No correlation was found between the size of Golgi complexes and cell area. The largest Golgi complexes were observed in the smallest cells.

The size of the Golgi complexes was not correlated with the cellular size. PECAM and ecNOS labeling revealed a considerable variation in endocardial EC size between various areas of the right and left ventricular endocardium. Since cellular borders could not be detected in arterial ECs after ecNOS staining, the reciprocal of cell density was used to estimate the area of ECs. Cellular area was analyzed in a manner similar to that used for the Golgi complex area. The coefficients of variation were 16.1%, 10.5%, and 16.8%, and cellular area was estimated to be 666±40 (mean±SE), 350±28, and 439±7 µm² for endocardial, arterial, and venous ECs, respectively. These estimated values were of the same magnitude as measurements of the cellular area from rat endothelium, where cellular borders were visualized after staining for actin or PECAM (References 32 and 34^{32 34} , and authors' unpublished data, 1996). Fig 5C confirms the lack of correlation between the Golgi complex area and the cellular area in rat cardiac endothelium. The largest Golgi complexes were present in arterial ECs, which had the smallest cellular area.

The shape of the Golgi complexes was nonuniformly distributed in cardiac endothelium. For the maximal cord of Golgi complexes, the coefficient of variation in arterial ECs, 31.8%, was significantly larger than that for endocardial or venous ECs, 23.7% and 26.9%, respectively. The maximal cord in endocardial ECs was significantly different from that in arterial or venous ECs (Fig 5B).

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
The present study demonstrates that all cardiac ECs expressed ecNOS but that there existed a considerable nonuniformity of ecNOS labeling between endocardial, arterial, capillary, and venous ECs. The more intense ecNOS staining in endocardial ECs and arterial endothelium appeared to be correlated with a more intensely labeled and larger-sized Golgi complex. Heterogeneity was also characteristic for ecNOS labeling of the peripheral cell borders. The ecNOS-stained peripheral borders were distinct in endocardial ECs, less distinct in venous ECs, nearly absent in arterial ECs, and not observed in capillary endothelium.

Our immunofluorescence data confirmed and extended previous observations of histochemical NOS detection in the heart. When NOS was determined by the histochemical NADPH-diaphorase method, it was demonstrated that coronary arteries in pig hearts manifested more NOS activity than did coronary veins.⁸ Weaker ecNOS immunoreactivity and NADPH-diaphorase staining of venous endothelium compared with arterial endothelium have also been described for lung and kidney.^{29 30} Moreover, stronger NADPH-diaphorase staining was present in the endothelium of efferent arterioles than in the endothelium of afferent arterioles of mammalian kidneys.³⁰ The strong NADPH-staining in ECs of efferent arterioles was ascribed to the mixed presence of ecNOS and nNOS (NOS type I).³⁰ The presence of the neuronal isoform of NOS has also been reported in endothelium from rabbit aorta and from rat coronary and pulmonary arteries.^{35 36 37} However, since macrophage-like cells were also labeled, the authors could not exclude the possibility that the nNOS antibody also reacted with other NOS isoforms.³⁷ Our present results with a commercial anti-nNOS polyclonal antibody did not provide evidence of nNOS expression in endocardial ECs and in the endothelium of coronary arteries of rat hearts. Further investigations are needed to support the absence or presence of nNOS in endocardial ECs or in the endothelium of coronary arteries.

Immunofluorescent staining of ecNOS strikingly labeled the Golgi complexes of ECs, as validated by double immunostaining for ecNOS and Golgi 58k protein. The subcellular distribution of ecNOS in Golgi complexes has also been documented in cultured endothelium using NADPH-diaphorase staining.³⁸ Spots of the histochemical reaction product have also been shown to be visible in situ, near nuclei in sections of endocardial endothelium (see Fig 2B of Reference 8⁸ ). Further biochemical analysis and oligonucleotide-directed mutagenesis determined that cotranslational N-myristoylation of ecNOS is necessary for ecNOS Golgi targeting^{13 15} and that posttranslational palmitoylation influences ecNOS targeting into caveolae.^{16 39 40 41} In cardiac endothelium, the largest Golgi complexes were present in arterial and endocardial ECs. The size of the Golgi complexes was not related to the surface area of the ECs. Large and flattened ECs might possess a flattened and hence apparently larger Golgi complex. However, the largest Golgi complexes were present in the arterial ECs that had the smallest surface area. The size of the Golgi complex is probably a marker of the synthetic activity of a cell. Coronary arterial ECs and endocardial ECs in the rat might have a higher synthetic activity than do capillary and venous ECs.

Previously, ultrastructural studies in teleosts demonstrated that endocardial ECs contained more ribosomes, endoplasmic reticulum, and larger Golgi complexes than did ECs from myocardial capillaries,⁴² suggesting that endocardial ECs are more involved in protein synthesis than are capillary ECs. Ultrastructural investigation of endocardial ECs in rats also demonstrated well-developed Golgi complexes with many juxtanuclear coated and uncoated vesicles.^{34 43} In ECs, the Golgi complex is involved in the synthesis of various proteins, ranging from extracellular matrix components like collagen⁴⁴ to more typical endothelial components like the intercellular adhesion molecule PECAM-1 (Fig 2A), the von Willebrand factor,^{45 46} and coagulation factor S.⁴⁷ Hypertrophied Golgi complexes and proliferation of endoplasmic reticulum in endothelium are characteristic for embryological processes,⁴⁸ for endothelial regeneration,⁴⁹ and in dysfunctional endothelium during various pathological conditions, such as hypercholesterolemia,⁵⁰ endotoxin injury,^{51 52} chronic ethanol administration,⁵³ and hydrostatic edema formation.⁵⁴ The size of the Golgi complex can thus be used as a marker for the functional status of ECs.

Intense labeling and the large size of Golgi complexes in endocardial ECs and in coronary arterial endothelium indicated a high rate of ecNOS production and could be related to the strong cytoplasmic ecNOS expression in these cells. Experiments in cultured ECs have demonstrated that ecNOS expression can be modulated by shear stress,²³ transforming growth factor-ß,²⁵ protein kinase C,⁵⁵ tumor necrosis factor-,²⁶ oxygen,²⁴ and the proliferative state.⁵⁶ In the heart, differences in shear stress could explain the strong and weak ecNOS expression of arterial and venous ECs, respectively. Experiments with various reporter systems and direct measurements of NO have demonstrated that laminar shear stress increases the endothelial release of NO.^{57 58 59} Fluid shear stress increases not only NOS mRNA⁵⁸ and protein but also endothelial superoxide dismutase, which further augments the local release of NO.⁶⁰ Shear stress might thus be involved in the differential expression of ecNOS in arterial, capillary, and venous ECs of rat hearts. But what about endocardial ECs? By comparison, laminar fluid shear stress is probably not high along the surface of endocardial ECs; nevertheless, although less pronounced than arterial ECs, endocardial ECs also manifested strong ecNOS expression. The endocardial surface might be more subjected to turbulent flow, yet this type of flow does not increase NOS mRNA and NO release in cultured human umbilical vein ECs.⁵⁸ In endocardial ECs, mechanical strain by three-dimensional changes of the inner wall during the cardiac cycle might influence ecNOS expression. ECs cultured on flexible substrates and subjected to cyclic strain showed an increase of NOS mRNA, protein, and NO production.⁶¹ However, we did not observe significant differences in ecNOS expression between various areas of the endocardial endothelium known to undergo distinct differences in mechanical deformation during the cardiac cycle, eg, the tendon end of right ventricular papillary muscles and the atrioventricular valves. Endocardial ECs covering these highly elastic structures are smaller and have a cytoskeletal organization^{34 62} different from that of other endocardial areas, but they did not show consistent differences in ecNOS labeling and in the size of Golgi complexes. Remarkably, freshly isolated ECs from large porcine coronary arteries do express more ecNOS protein and produce more NO than do ECs from resistance arterioles,⁶³ although both are subjected to similar shear stress. Factors other than shear stress might thus influence the expression of ecNOS in cardiac endothelium.

A distinct finding in endocardial ECs and in coronary venous ECs, but much less in coronary arterial endothelium and not in capillary endothelium, was the presence of bands of ecNOS labeling along the cell periphery. These bands coincided with the area of intercellular contacts, as demonstrated by double immunostaining with PECAM-1. PECAM-1 is an intercellular adhesion molecule that labels the whole depth of endothelial intercellular clefts.⁶⁴ Remarkably, a similar labeling of peripheral borders in endocardial ECs was present after NADPH-diaphorase histochemical staining (see Fig 2B in Reference 8⁸ ). This suggests enzymatic ecNOS activity along the peripheral borders of endocardial ECs. Previous studies involving cultured ECs have demonstrated that an NO-induced increase of cGMP decreases paracellular permeability.^{65 66 67} NO production by the peripherally located ecNOS in endocardial and venous ECs might thus be involved in the regulation of paracellular permeability.

Surprisingly, the ecNOS-rich intercellular boundaries of endocardial ECs were not stained or were only weakly stained for caveolin. Caveolin-rich microdomains in the plasmalemma are sites where ecNOS and various other molecules involved in transduction mechanisms are situated.²⁰ Depending on the type of the vascular bed, the number of caveolae can range from 10 to 460/µm² of EC surface.⁶⁸ Besides an association with plasma membrane caveolae, caveolin also resides in the Golgi complex and appears to cycle between these two compartments.^{69 70} Caveolin staining, in contrast to ecNOS labeling, was much stronger in the endothelium of myocardial capillaries than in endocardial and arterial ECs. This apparently confirms that compared with large vessel endothelium, microvascular endothelium contains more caveolae.^{40 71} Transmission electron microscopy revealed less caveolar vesicles and pits in endocardial ECs than in the endothelium of myocardial capillaries.³⁴ Further work is clearly needed to determine the correlation between the presence of caveolin and ecNOS in the various vascular beds. The distinct ecNOS labeling and absence of caveolin labeling of peripheral borders in endocardial ECs suggest that ecNOS might also be associated with other membrane components or with cytoskeletal components.

The distinct ecNOS labeling of Golgi complex and peripheral borders in endocardial ECs raises the question of where the active pool of ecNOS is located. Our results cannot address the existence of enzymatic ecNOS activity in a particular compartment. The enzymatic activity of Golgi complexes is probably reflected by its NADPH-diaphorase activity and staining of ECs in vitro³⁸ and in situ.^{8 17} Fractionating studies suggest the presence of mature ecNOS in EC membranes and in intracellular membranes.⁴⁰ Cytosolic fractions contain a substantially lower ecNOS activity than does the particulate fraction.⁷² Brefeldin-induced disassembly of the Golgi complex in cultured ECs results in a loss of NADPH-diaphorase activity and in a significant decrease of NO production.⁷³ This decrease was already significant at 15 minutes and was maximal at 90 minutes after incubation with brefeldin. This decrease is probably not an effect of a blockade of ecNOS cycling from Golgi complex to the plasma membrane, since the half-life of ecNOS protein⁴¹ measures 20 hours. The mechanisms of the brefeldin-induced inhibition of NO production are not known. The inhibition might result indirectly from interference with a cofactor needed for enzyme activity or from blocking of the active site.⁷³ However, these data probably support the hypothesis that Golgi complexes, besides the caveolar membrane domains, are important sites of NO production.

In conclusion, the present study demonstrated considerable nonuniformity in the expression of ecNOS and of the size of Golgi complexes in cardiac endothelium. The presence of intense ecNOS-labeled and large-sized Golgi complexes in endocardial ECs and coronary arterial cells is in accordance with a more intense cytoplasmic ecNOS labeling and is suggestive for a higher ecNOS activity in these cells than in coronary venous and myocardial capillary ECs. The lack of caveolin labeling and the presence of ecNOS labeling along the periphery of endocardial and venous ECs suggest that ecNOS might be associated with other membrane components or with parts of the cytoskeleton. Further investigations in disease states or during embryological development might allow a better understanding of ecNOS distribution and the size of Golgi complexes in cardiac endothelium.

	Selected Abbreviations and Acronyms

 

DTAF = dichlorotriazinylamino fluorescein EC = endothelial cell ecNOS = endothelial constitutive NOS nNOS = neuronal NOS NOS = NO synthase PECAM = platelet and EC adhesion molecule RECA = rat EC antibody

	Acknowledgments

 
This study was supported by a grant from the Belgian Program of Interuniversitary Poles of Attraction, initiated by the Belgian State, Prime Ministers Office, Science Policy Programming.

Received September 8, 1997; accepted October 1, 1997.

	References

Top Abstract Introduction Materials and Methods Results Discussion References

 

1. Finkel MS, Oddis CV, Jacob TD, Watkins SC, Hattler BG, and Simmons RL. Negative inotropic effects of cytokines on the heart mediated by nitric oxide. Science. 1992;257:387–389.[Abstract/Free Full Text]
2. Li K, Sirois P, and Rouleau JL. Role of endothelial cells in cardiovascular function. Life Sci. 1994;54:579–592.[Medline] [Order article via Infotrieve]
3. Shah AM. Paracrine modulation of heart cell function by endothelial cells. Cardiovasc Res. 1996;31:847–867.[Medline] [Order article via Infotrieve]
4. Belhassen L, Kelly RA, Smith TW, Balligand JL. Nitric oxide synthase (NOS3) and contractile responsiveness to adrenergic and cholinergic agonists in the heart: regulation of NOS3 transcription in vitro and in vivo by cyclic adenosine monophosphate in rat cardiac myocytes. J Clin Invest. 1996;97:1908–1915.[Medline] [Order article via Infotrieve]
5. Mohan P, Brutsaert DL, Paulus WJ, Sys SU. Myocardial contractile response to nitric oxide and cGMP. Circulation. 1996;93:1223–1229.[Abstract/Free Full Text]
6. Paulus WJ, Vantrimpont PJ, Shah AM. Paracrine coronary endothelial control of left ventricular function in humans. Circulation. 1995;92:2119–2126.[Abstract/Free Full Text]
7. Tanaka K, Hassall CJ, Burnstock G. Distribution of intracardiac neurones and nerve terminals that contain a marker for nitric oxide, NADPH-diaphorase, in the guinea-pig heart. Cell Tissue Res. 1993;273:293–300.[Medline] [Order article via Infotrieve]
8. Ursell PC, and Mayes M. The majority of nitric oxide synthase in pig heart is vascular and not neural. Cardiovasc Res. 1993;27:1920–1924.[Abstract/Free Full Text]
9. Ursell PC, Mayes M. Anatomic distribution of nitric oxide synthase in the heart. Int J Cardiol. 1995;50:217–223.[Medline] [Order article via Infotrieve]
10. Wildhirt SM, Dudek RR, Suzuki H, Pinto V, Narayan KS, Bing RJ. Immunohistochemistry in the identification of nitric oxide synthase isoenzymes in myocardial infarction. Cardiovasc Res. 1995;29:526–531.[Medline] [Order article via Infotrieve]
11. Feron O, Belhassen L, Kobzik L, Smith TW, Kelly RA, Michel T. Endothelial nitric oxide synthase targeting to caveolae:. specific interactions with caveolin isoforms in cardiac myocytes and endothelial cells. J Biol Chem. 1996;271:22810–22814.[Abstract/Free Full Text]
12. Busconi L, Michel T. Endothelial nitric oxide synthase: N-terminal myristoylation determines subcellular localization. J Biol Chem. 1993;268:8410–8413.[Abstract/Free Full Text]
13. Sessa WC, Barber CM, Lynch KR. Mutation of N-myristoylation site converts endothelial cell nitric oxide synthase from a membrane to a cytosolic protein. Circ Res. 1993;72:921–924.[Abstract/Free Full Text]
14. Hecker M, Mulsch A, Bassenge E, Forstermann U, Busse R. Subcellular localization and characterization of nitric oxide synthase(s) in endothelial cells: physiological implications. Biochem J. 1994;299:247–252.
15. Sessa WC, Garcia-Cardena G, Liu J, Keh A, Pollock JS, Bradley J, Thiru S, Braverman IM, Desai KM. The Golgi association of endothelial nitric oxide synthase is necessary for the efficient synthesis of nitric oxide. J Biol Chem. 1995;270:17641–17644.[Abstract/Free Full Text]
16. Shaul PW, Smart EJ, Robinson LJ, German Z, Yuhanna IS, Ying YS, Anderson RGW, Michel T. Acylation targets endothelial nitric-oxide synthase to plasmalemmal caveolae. J Biol Chem. 1996;271:6518–6522.[Abstract/Free Full Text]
17. O'Brien AJ, Young HM, Povey JM, Furness JB. Nitric oxide synthase is localized predominantly in the Golgi apparatus and cytoplasmic vesicles of vascular endothelial cells. Histochem Cell Biol. 1995;103:221–225.[Medline] [Order article via Infotrieve]
18. Mellman I, Simons K. The Golgi complex: in vitro veritas? Cell. 1992;68:829–840.[Medline] [Order article via Infotrieve]
19. Rothberg KG, Heuser JE, Donzell WC, Ying YS, Glenney JR, Anderson RG. Caveolin, a protein component of caveolae membrane coats. Cell. 1992;68:673–682.[Medline] [Order article via Infotrieve]
20. Schnitzer JE, Mcintosh DP, Dvorak AM, Liu J, Oh P. Separation of caveolae from associated microdomains of GPI-anchored proteins. Science. 1995;269:1435–1439.[Abstract/Free Full Text]
21. Schnitzer JE, Oh P, Jacobson BS, Dvorak AM. Caveolae from luminal plasmalemma of rat lung endothelium: microdomains enriched in caveolin, Ca(2+)-ATPase, and inositol trisphosphate receptor. Proc Natl Acad Sci U S A. 1995;92:1759–1763.[Abstract/Free Full Text]
22. Garcia-Cardena G, Fan R, Stern DF, Liu J, Sessa WC. Endothelial nitric oxide synthase is regulated by tyrosine phosphorylation and interacts with caveolin-1. J Biol Chem. 1996;271:27237–27240.[Abstract/Free Full Text]
23. Uematsu M, Ohara Y, Navas JP, Nishida K, Murphy TJ, Alexander RW, Nerem RM, Harrison DG. Regulation of endothelial cell nitric oxide synthase mRNA expression by shear stress. Am J Physiol. 1995;269:C1371–C1378.[Abstract/Free Full Text]
24. North AJ, Lau KS, Brannon TS, Wu LJC, Wells LB, German Z, Shaul PW. Oxygen upregulates nitric oxide synthase gene expression in ovine fetal pulmonary artery endothelial cells. Am J Physiol. 1996;14:L643–L649.
25. Inoue N, Venema RC, Sayegh HS, Ohara Y, Murphy TJ, Harrison DG. Molecular regulation of the bovine endothelial cell nitric oxide synthase by transforming growth factor-beta 1. Arterioscler Thromb Vasc Biol. 1995;15:1255–1261.[Abstract/Free Full Text]
26. Yoshizumi M, Perrella MA, Burnett JC Jr, Lee ME. Tumor necrosis factor downregulates an endothelial nitric oxide synthase mRNA by shortening its half-life. Circ Res. 1993;73:205–209.[Abstract]
27. Harrison DG, Sayegh H, Ohara Y, Inoue N, Venema RC. Regulation of expression of the endothelial cell nitric oxide synthase. Clin Exp Pharmacol Physiol. 1996;23:251–255.[Medline] [Order article via Infotrieve]
28. Nava E, Noll G, Luscher TF. Nitric oxide in cardiovascular diseases. Ann Med. 1995;27:343–351.[Medline] [Order article via Infotrieve]
29. Pollock JS, Nakane M, Buttery LD, Martinez A, Springall D, Polak JM, Forstermann U, Murad F. Characterization and localization of endothelial nitric oxide synthase using specific monoclonal antibodies. Am J Physiol. 1993;265:C1379–C1387.[Abstract/Free Full Text]
30. Bachmann S, Bosse HM, Mundel P. Topography of nitric oxide synthesis by localizing constitutive NO synthases in mammalian kidney. Am J Physiol. 1995;268:F885–F898.[Abstract/Free Full Text]
31. Tang BL, Low SH, Hong WJ. Differential response of resident proteins and cycling proteins of the Golgi to brefeldin A. Eur J Cell Biol. 1995;68:199–205.[Medline] [Order article via Infotrieve]
32. Andries LJ, Brutsaert DL. Endocardial endothelium: junctional organization and permeability. Cell Tissue Res. 1994;277:391–400.[Medline] [Order article via Infotrieve]
33. Duijvestijn AM, Vangoor H, Klatter F, Majoor GD, Vanbussel, Vriesman PJCV. Antibodies defining rat endothelial cells: RECA-1, a pan-endothelial cell-specific monoclonal antibody. Lab Invest. 1992;66:459–466.[Medline] [Order article via Infotrieve]
34. Andries LJ. Endocardial Endothelium: Functional Morphology. Austin, Tex: RG Landes Co; 1994.
35. Dikranian K, Trosheva M, Nikolov S, Bodin P. Nitric oxide synthase (NOS) in the human umbilical cord vessels: an immunohistochemical study. Acta Histochem. 1994;96:145–153.[Medline] [Order article via Infotrieve]
36. Loesch A, Belai A, Burnstock G. Ultrastructural localization of NADPH-diaphorase and colocalization of nitric oxide synthase in endothelial cells of the rabbit aorta. Cell Tissue Res. 1993;274:539–545.[Medline] [Order article via Infotrieve]
37. Loesch A, Burnstock G. Ultrastructural localization of nitric oxide synthase and endothelin in rat pulmonary artery and vein during postnatal development and ageing. Cell Tissue Res. 1996;283:355–365.[Medline] [Order article via Infotrieve]
38. Morin AM, Stanboli A. Nitric oxide synthase localization in cultured cerebrovascular endothelium during mitosis. Exp Cell Res. 1994;211:183–188.[Medline] [Order article via Infotrieve]
39. Liu J, Garcia-Cardena G, Sessa WC. Palmitoylation of endothelial nitric oxide synthase is necessary for optimal stimulated release of nitric oxide: implications for caveolae localization. Biochemistry. 1996;35:13277–13281.[Medline] [Order article via Infotrieve]
40. Garcia-Cardena G, Oh P, Liu J, Schnitzer JE, Sessa WC. Targeting of nitric oxide synthase to endothelial cell caveolae via palmitoylation: implications for nitric oxide signaling. Proc Natl Acad Sci U S A. 1996;93:6448–6453.[Abstract/Free Full Text]
41. Robinson LJ, Busconi L, Michel T. Agonist-modulated palmitoylation of endothelial nitric oxide synthase. J Biol Chem. 1995;270:995–998.[Abstract/Free Full Text]
42. Midttun B. Ultrastructure of atrial and ventricular endocardium and cardiac capillary endothelium of the pike. Tissue Cell. 1981;13:747–756.[Medline] [Order article via Infotrieve]
43. Brutsaert DL, Andries LJ. The endocardial endothelium. Am J Physiol. 1992;263:H985–H1002.[Abstract/Free Full Text]
44. Hayashi M, Ninomya Y, Hayashi K, Linsenmayer TF, Olsen BR, Trelstad RL. Secretion of collagen types I and II by epithelial and endothelial cells in the developing chick cornea demonstrated by in situ hybridization and immunohistochemistry. Development. 1988;103:27–36.[Abstract]
45. Sinha S, Wagner DD. Intact microtubules are necessary for complete processing, storage and regulated secretion of von Willebrand factor by endothelial cells. Eur J Cell Biol. 1987;43:377–383.[Medline] [Order article via Infotrieve]
46. Jeanneau C, Sultan Y. Localization of factor VIII/von Willebrand factor antigen by immunoelectron microscopy in human endothelial cells using Fab fragments coupled to peroxidase. J Histochem Cytochem. 1982;30:1091–1096.[Abstract]
47. Stern D, Brett J, Harris K, Nawroth P. Participation of endothelial cells in the protein C-protein S anticoagulant pathway: the synthesis and release of protein S. J Cell Biol. 1986;102:1971–1978.[Abstract/Free Full Text]
48. Markwald RR, Fitzharris TP, Adams Smith WN. Structural analysis of endocardial cytodifferentiation. Dev Biol. 1975;42:160–180.[Medline] [Order article via Infotrieve]
49. Turcotte H, Bazin M, Boutet M. Junctional complexes in regenerating endocardium. J Ultrastruct Res. 1982;79:133–141.[Medline] [Order article via Infotrieve]
50. Takasu N, Kurihara H, Uchida K. Early changes in fine structures of the aortic arch in hamsters fed a high cholesterol diet. Arch Histol Cytol. 1990;53:211–218.[Medline] [Order article via Infotrieve]
51. Reidy MA, Chopek M, Chao S, McDonald T, Schwarz SM. Injury induces increase of von Willebrand factor in rat endothelial cells. Am J Pathol. 1989;134:857–864.[Abstract]
52. Freudenberg N, Madreiter H, Mittermayer C. Experimental investigations into the pathogenesis of endocarditis due to shock. Beitr Pathol. 1975;155:248–262.[Medline] [Order article via Infotrieve]
53. Karwacka H. Ultrastructural and biochemical studies of the brain and other organs in rats after chronic ethanol administration. Exp Pathol. 1980;18:118–126.
54. DeFouw DO. Morphologic study of the alveolar septa in normal and edematous isolated dog lungs fixed by vascular perfusion. Lab Invest. 1980;42:413–419.[Medline] [Order article via Infotrieve]
55. Ohara Y, Sayegh HS, Yamin JJ, Harrison DG. Regulation of endothelial constitutive nitric oxide synthase by protein kinase C. Hypertension. 1995;25:415–420.[Abstract/Free Full Text]
56. Arnal JF, Yamin J, Dockery S, Harrison DG. Regulation of endothelial nitric oxide synthase mRNA, protein, and activity during cell growth. Am J Physiol. 1994;267:C1381–C1388.[Abstract/Free Full Text]
57. Lamontagne D, Pohl U, Busse R. Mechanical deformation of vessel wall and shear stress determine the basal release of endothelium-derived relaxing factor in the intact rabbit coronary vascular bed. Circ Res. 1992;70:123–130.[Abstract/Free Full Text]
58. Noris M, Morigi M. Nitric oxide synthesis by cultured endothelial cells is modulated by flow conditions. Circ Res. 1995;76:536–543.[Abstract/Free Full Text]
59. Kanai AJ, Strauss HC, Truskey GA, Crews AL, Grunfeld S, Malinski T. Shear stress induces ATP-independent transient nitric oxide release from vascular endothelial cells measured directly with a porphyrinic microsensor. Circ Res. 1995;77:284–293.[Abstract/Free Full Text]
60. Inoue N, Ramasamy S, Fukai T, Nerem RM, Harrison DG. Shear stress modulates expression of Cu/Zn superoxide dismutase in human aortic endothelial gels. Circ Res. 1996;79:32–37.[Abstract/Free Full Text]
61. Awolesi MA, Sessa WC, and Sumpio BE. Cyclic strain upregulates nitric oxide synthase in cultured bovine aortic endothelial cells. J Clin Invest. 1995;96:1449–1454.
62. Andries LJ, Brutsaert DL. Endocardial endothelium in rat heart: cell shape and organization of the cytoskeleton. Cell Tissue Res. 1993;273:107–117.[Medline] [Order article via Infotrieve]
63. Xu XP, Liu Y, Tanner MA, Sturek M, Myers PR. Differences in nitric oxide production in porcine resistance and epicardial conduit coronary arteries. J Cell Physiol. 1996;168:539–548.[Medline] [Order article via Infotrieve]
64. Leach L, Clark P, Lampugnani MG, Arroyo AG, Dejana E, Firth JA. Immunoelectron characterisation of the inter-endothelial junctions of human term placenta. J Cell Sci. 1993;104:1073–1081.[Abstract]
65. Westendorp RG, Draijer R, Meinders AE, van Hinsbergh VW. Cyclic-GMP-mediated decrease in permeability of human umbilical and pulmonary artery endothelial cell monolayers. J Vasc Res. 1994;31:42–51.[Medline] [Order article via Infotrieve]
66. Vigne P, Lund L, Frelin-C. Cross talk among cyclic AMP, cyclic GMP, and Ca(2+)-dependent intracellular signalling mechanisms in brain capillary endothelial cells. J Neurochem. 1994;62:2269–2274.[Medline] [Order article via Infotrieve]
67. Oliver JA. Endothelium-derived relaxing factor contributes to the regulation of endothelial permeability. J Cell Physiol. 1992;151:506–511.[Medline] [Order article via Infotrieve]
68. Wagner RC, Casley-Smith JR. Endothelial vesicles. Microvasc Res. 1981;21:267–298.[Medline] [Order article via Infotrieve]
69. Conrad PA, Smart EJ, Ying YS, Anderson RGW, Bloom GS. Caveolin cycles between plasma membrane caveolae and the Golgi complex by microtubule-dependent and microtubule-independent steps. J Cell Biol. 1995;131:1421–1433.[Abstract/Free Full Text]
70. Smart EJ, Ying YS, Conrad PA, Anderson RG. Caveolin moves from caveolae to the Golgi apparatus in response to cholesterol oxidation. J Cell Biol. 1994;127:1185–1197.[Abstract/Free Full Text]
71. Kumar S, West DC, Ager A. Heterogeneity in endothelial cells from large vessels and microvessels. Differentiation. 1987;36:57–70.[Medline] [Order article via Infotrieve]
72. Mitchell JA, Forstermann U, Warner TD, Pollock JS, Schmidt HH, Heller M, Murad F. Endothelial cells have a particulate enzyme system responsible for EDRF formation: measurement by vascular relaxation. Biochem Biophys Res Commun. 1991;176:1417–1423.[Medline] [Order article via Infotrieve]
73. Stanboli A, Morin AM. Nitric oxide synthase in cerebrovascular endothelial cells is inhibited by brefeldin A. Neurosci Lett. 1994;171:209–212.[Medline] [Order article via Infotrieve]