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Endothelial Cell Regulation of Pulmonary Vascular Tone, Inflammation, and Coagulation

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ABSTRACT

The pulmonary endothelium represents a heterogeneous cell monolayer covering the luminal surface of the entire lung vasculature. As such, this cell layer lies at a critical interface between the blood, airways, and lung parenchyma, and must act as a selective barrier between these diverse compartments. Lung endothelial cells are able to produce and secrete mediators, display surface receptor, and cellular adhesion molecules, and metabolize circulating hormones to influence vasomotor tone, both local and systemic inflammation, and coagulation functions. In this review, we will explore the role of the pulmonary endothelium in each of these systems, highlighting key regulatory functions of the pulmonary endothelial cell, as well as novel aspects of the pulmonary endothelium in contrast to the systemic cell type. The interactions between pulmonary endothelial cells and both leukocytes and platelets will be discussed in detail, and wherever possible, elements of endothelial control over physiological and pathophysiological processes will be examined. © 2015 American Physiological Society. Compr Physiol 5:531‐559, 2015.

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Figure 1. Figure 1. Endothelial control of pulmonary vascular tone. An increase in intracellular Ca2+, either from internal stores or the extracellular fluid, triggers a series of intracellular signaling events that culminate in the binding of Ca2+ to calmodulin, activation of myosin light chain kinase (MLCK), and the phosphorylation of myosin. This, in turn, causes cell contraction and vasoconstriction. The most important regulator of this process in nitric oxide (NO). NO activates soluble guanylate cyclase (sGC), which produces cGMP. cGMP, acting via protein kinase G (PKG), can then inhibit various membrane channels to block the vasoconstrictive response. Additionally, PKG can inhibit sarcoplasmic reticulum (SR) Ca2+ channels, and can release myosin light chain phosphatase (MLCP) from inhibition by ROK, leading to dephosphorylation of MLC and the cessation of SMC contraction.
Figure 2. Figure 2. The pulmonary endothelial cell in inflammation. As highlighted in the text, the interaction between the pulmonary endothelium and circulating leukocytes differs greatly from that in the systemic circulation. Important physical factors exist which lead to leukocyte retention in the pulmonary capillary network. To traverse the lung vasculature, neutrophils must undergo cytoskeletal rearrangement so that they can fit through the small pulmonary vessels, which also have far lower flow rates than the systemic circulation. Because of this, the role of leukocyte rolling is diminished in the lung, and both CD18‐dependent and CD18‐independnet pathways exist for leukocyte extravasation.
Figure 3. Figure 3. The role of the pulmonary endothelium in platelet function. At rest, the pulmonary EC represents an antithrombogenic surface, repelling interactions with both platelets and clotting factors. This antiplatelet phenotype is largely due to constitutive secretion of PGI2 and NO by the resting lung EC. After endothelial injury, or inflammatory states of platelet and EC activation, the EC will secrete proplatelet aggregation factors, such as thromboxane A2. Activated platelets have increased availability of GPIIb/IIIa, which binds to elevated surface levels of ICAM‐1 and integrins. Furthermore, endothelial‐derived von Willebrand Factor (VWF) is released, and assists in cross‐linking platelets to form a mature platelet plug.
Figure 4. Figure 4. Regulation of nitric oxide production in the pulmonary endothelium. eNOS forms NO in the lung EC from L‐arginine, in the presence of the cofactor BH4, as a byproduct of L‐citrulline production. eNOS activity is enhanced by shear stress, intracellular Ca2+, and signaling by cytokines such as TGF‐β. Calcium enhances eNOS activity via direct binding of Ca2+/calmodulin (CaM) to eNOS, while phosphorylation of eNOS at Ser1177 by PI3‐Kinase and Akt also increases NO production. NO produced in this manner can freely diffuse into the PASMC, where it activates soluble guanylate cyclase to produce cGMP. cGMP then, via PKG function, produces PASMC relaxation. In the EC itself, NO inhibits neutrophil and platelet binding. On the other hand, reactive oxygen species (ROS), and conditions favoring uncoupling (including loss of BH4 availability and inflammatory states) decrease eNOS activity and NO production. Binding of eNOS to caveolin‐1 in caveolae downregulates is activity, as does tyrosine phosphorylation by PKC. Under such conditions, in addition to diminished NO production, uncoupled eNOS will produce superoxide, which then leads to further uncoupling in a positive feedback loop.
Figure 5. Figure 5. Arachidonic acid metabolism. The enzyme cytosolic phospholipase A2 (cPLA2), which is the dominant isoform in the pulmonary EC, liberates the 20 carbon fatty acid, arachidonic acid (AA) from plasma membrane phospholipids. Among several possible fates for AA, it can be metabolized by lipooxygenases (LOX) to produce Hydroperoxyeicosatetraenoic acid (HPETE), which can be further metabolized into leukotrienes. The activity of cyclooxygenase enzymes (COX) on AA produces prostaglandin H2 (PGH2) which can then be further processed into one of PGI2, PGE2, or thromboxane A2 (TxA2).
Figure 6. Figure 6. Pulmonary endothelial synthesis of vasoactive EET and HETE species. Further pathways of AA metabolism in the pulmonary endothelial cell can produce EET and HETE. The action of the cytochrome P450 epoxygenases 2C and 2J can produce a family of EET species, with various vasomotor and inflammatory functions, detailed in the text. The main catabolizer of EET species is the soluble epoxide hydrolase (sEH), which breaks EET species down into dihydroxyeicosatrienoic acids (DHET). Alternatively, ω or ω‐1 hydrolases can metabolize AA into 20‐ or 19‐HETE.
Figure 7. Figure 7. Pulmonary endothelial synthesis and secretion of ET‐1. ET‐1 is produced in the pulmonary endothelial cell in a prepro form, which is cleaved into pro‐ET‐1 (also called big ET‐1). The enzyme endothelin converting enzyme (ECE) then cleaves big ET‐1 into mature ET‐1, which then has both autocrine and paracrine functions within the lung vasculature. In the EC itself, ET‐1 can bind to the ETB receptor (ETBR), which results in nitric oxide (NO) and prostacyclin (PGI2) production. These two mediators can both subsequently act upon the smooth muscle cell to cause vasorelaxation. In a paracrine fashion, ET‐1 itself can bind to its receptors on smooth muscle cells, ETAR and ETBR, to cause G‐protein‐mediated activation of phospholipase C (PLC). The resultant production of inositol triphosphate (IP3) and diacylglycerol (DAG) results in SMC migration, and contraction, or proliferation, respectively.
Figure 8. Figure 8. Sources of reactive oxygen species in the pulmonary endothelial cell. A critically important source of ROS in the pulmonary EC is the family of NADPH oxidase (NOX) enzymes. Activated by angiotensin II, TGF‐β, and shear stress, NOX isoforms present in the lung can generate large quantities of ROS. Similarly, the mitochondrial electron transport chain can produce ROS, although controversy exists regarding whether hyperoxia or hypoxia favors this condition, and this controversy is discussed in the main text. Importantly, uncoupled eNOS produces ROS, and this ROS then leads to further eNOS uncoupling, which is an important propagator of inflammatory stress under a variety of pathological states. Elevated ROS levels leads to endothelial cytoskeletal changes, increased surface adhesion molecule expression, platelet adhesion, and a state referred to as endothelial dysfunction. Important enzymes such as superoxide dismutase exist to metabolize ROS and limit its accumulation.
Figure 9. Figure 9. Heterogeneous distribution of Weibel‐Pallade Bodies (WPB) in the lung endothelium. Representative electron micrographs of pulmonary endothelial cells from human lungs. WPB are indicated by arrowheads. Upper panels: High and low magnification images of extra‐alveolar endothelial cells, showing readily detectable WPB. Lower panels depict alveolar capillary endothelial cells, which are notably devoid of WPB. EC, endothelial cells, N, nucleus, BM, basement membrane, *, caveolae. Reproduced, with permission, from ().
Figure 10. Figure 10. Transcytosis of interleukin‐8 by endothelial cells. Stacked and reconstructed electron micrographs of human skin venular endothelial cells, with gold labeling of interleukin‐8 (IL‐8). (A) IL‐8 binding and uptake at the basolateral (abluminal) surface of the venular EC. (B) and (C) demonstrate IL‐8 in vesicles below the plasma membrane. (D) and (E) demonstrate IL‐8 now subsequently appearing at the luminal surface of the EC. (F) Colocalization of IL‐8 (10 nm gold) and caveolin‐1 (5 nm gold). Reproduced, with permission, from ().
Figure 11. Figure 11. Structure of the endothelial glycocalyx and endothelial surface layer (ESL). At the luminal surface of the pulmonary EC, the glycocalyx comprises a thin layer of plasma membrane glycoproteins and glycolipids. External to that is a thicker layer of glycosaminoglycans (GAG), heparan sulfate, hyaluronan, and other molecules, termed the ESL. These layers (A) separate the EC surface from the circulating plasma, (B) allow for mechanosensation and outside‐in signaling to the EC, (C) performs a critical role in maintaining the endothelial barrier, and (D) can be importantly degraded and modified during periods of inflammation.
Figure 12. Figure 12. The endothelial glycocalyx in vivo, and its degradation by LPS. (A) Schematic representation of the glycocalyx and endothelial surface layer (ESL). Note that the glycocalyx is not permeable to large molecules such as fluorescent dextrans. Therefore, measurement of vessel diameter by visible light microscopy versus fluorescence should allow for the measurement of the ESL. The images in the right of the panel show mouse sub‐pleural microvessels (MV) imaged with visible light (DIC) or fluorescence (FITC). The difference between the DIC and FITC width represents the ESL. (A) Alveolus. (B) Group data of calculated ESL thickness as in (A), taken from mice treated with LPS or TNF‐α, demonstrating how inflammatory stimuli can degrade the ESL. (C) These effects are not seen in mice lacking functional TNF receptors. Reproduced, with permission, from ().
Figure 13. Figure 13. Pulmonary endothelial angiotensin signaling. The majority of conversion of AngI to AngII is carried out by angiotensin converting enzyme (ACE), bound to the EC surface. AngII can then bind to AT1R or AT2R (left side). Signaling through AT1R results in G‐protein‐mediated release of IP3 and DAG, as well as stimulation of MAP kinases. The downstream effects include PASMC contraction and migration, as well as EC ROS production and vascular remodeling. Signaling via the AT2R is incompletely understood in the lung, but in general antagonizes the effects of AT1R signaling, leading to PASMC relaxation, and protection from adverse remodeling. The cell on the right depicts further metabolism of AngII by ACE2, producing Ang(1‐7), which subsequently binds the Mas receptor. Mas has a highly protective effect during states of acute lung injury and inflammation, decreasing ROS production, enhancing NO release, and protecting against thrombosis and EC apoptosis.
Figure 14. Figure 14. The conducted response in hypoxic pulmonary vasoconstriction. (A) Intravital microscopic images taken from wild‐type (top panels) or Cx40−/− mice (bottom panels) during normoxic or hypoxic ventilation, as indicated. Endothelial cells have been loaded with the voltage‐sensitive fluorophore, di‐8‐ANEPPS. Arteriolar vessel margins are indicated by dotted lines. Capillary endothelial cells are circled, while arteriolar EC are indicated by squares. Not that in wild‐type mice, depolarization in response to hypoxia occurs in both the capillary and arteriolar EC. However, in Cx40−/− mice, only the capillary EC depolarize, indicating the requirement of gap junction communication for the conducted response. (C) and (D) show group data describing the above. In (D), depolarization in response to hypoxia is not seen in the pulmonary arteriolar EC in Cx40−/− mice, indicative of the requirement for gap junctional communication. Reproduced, with permission, from ().
Figure 15. Figure 15. Schematic diagram representing the conducted response. It is hypothesized that hypoxia inhibits oxygen‐sensitive K+ channels, leading to EC depolarization at the alveolar capillary level. This depolarization spreads between EC in a retrograde fashion to the muscularized arteriolar EC via Cx40‐containing gap junctions. At this point, communication with the PASMC can occur, resulting in vasoconstriction. A contribution of myoendothelial gap junctions containing Cx43 and Cx40 is likely involved in this process.


Figure 1. Endothelial control of pulmonary vascular tone. An increase in intracellular Ca2+, either from internal stores or the extracellular fluid, triggers a series of intracellular signaling events that culminate in the binding of Ca2+ to calmodulin, activation of myosin light chain kinase (MLCK), and the phosphorylation of myosin. This, in turn, causes cell contraction and vasoconstriction. The most important regulator of this process in nitric oxide (NO). NO activates soluble guanylate cyclase (sGC), which produces cGMP. cGMP, acting via protein kinase G (PKG), can then inhibit various membrane channels to block the vasoconstrictive response. Additionally, PKG can inhibit sarcoplasmic reticulum (SR) Ca2+ channels, and can release myosin light chain phosphatase (MLCP) from inhibition by ROK, leading to dephosphorylation of MLC and the cessation of SMC contraction.


Figure 2. The pulmonary endothelial cell in inflammation. As highlighted in the text, the interaction between the pulmonary endothelium and circulating leukocytes differs greatly from that in the systemic circulation. Important physical factors exist which lead to leukocyte retention in the pulmonary capillary network. To traverse the lung vasculature, neutrophils must undergo cytoskeletal rearrangement so that they can fit through the small pulmonary vessels, which also have far lower flow rates than the systemic circulation. Because of this, the role of leukocyte rolling is diminished in the lung, and both CD18‐dependent and CD18‐independnet pathways exist for leukocyte extravasation.


Figure 3. The role of the pulmonary endothelium in platelet function. At rest, the pulmonary EC represents an antithrombogenic surface, repelling interactions with both platelets and clotting factors. This antiplatelet phenotype is largely due to constitutive secretion of PGI2 and NO by the resting lung EC. After endothelial injury, or inflammatory states of platelet and EC activation, the EC will secrete proplatelet aggregation factors, such as thromboxane A2. Activated platelets have increased availability of GPIIb/IIIa, which binds to elevated surface levels of ICAM‐1 and integrins. Furthermore, endothelial‐derived von Willebrand Factor (VWF) is released, and assists in cross‐linking platelets to form a mature platelet plug.


Figure 4. Regulation of nitric oxide production in the pulmonary endothelium. eNOS forms NO in the lung EC from L‐arginine, in the presence of the cofactor BH4, as a byproduct of L‐citrulline production. eNOS activity is enhanced by shear stress, intracellular Ca2+, and signaling by cytokines such as TGF‐β. Calcium enhances eNOS activity via direct binding of Ca2+/calmodulin (CaM) to eNOS, while phosphorylation of eNOS at Ser1177 by PI3‐Kinase and Akt also increases NO production. NO produced in this manner can freely diffuse into the PASMC, where it activates soluble guanylate cyclase to produce cGMP. cGMP then, via PKG function, produces PASMC relaxation. In the EC itself, NO inhibits neutrophil and platelet binding. On the other hand, reactive oxygen species (ROS), and conditions favoring uncoupling (including loss of BH4 availability and inflammatory states) decrease eNOS activity and NO production. Binding of eNOS to caveolin‐1 in caveolae downregulates is activity, as does tyrosine phosphorylation by PKC. Under such conditions, in addition to diminished NO production, uncoupled eNOS will produce superoxide, which then leads to further uncoupling in a positive feedback loop.


Figure 5. Arachidonic acid metabolism. The enzyme cytosolic phospholipase A2 (cPLA2), which is the dominant isoform in the pulmonary EC, liberates the 20 carbon fatty acid, arachidonic acid (AA) from plasma membrane phospholipids. Among several possible fates for AA, it can be metabolized by lipooxygenases (LOX) to produce Hydroperoxyeicosatetraenoic acid (HPETE), which can be further metabolized into leukotrienes. The activity of cyclooxygenase enzymes (COX) on AA produces prostaglandin H2 (PGH2) which can then be further processed into one of PGI2, PGE2, or thromboxane A2 (TxA2).


Figure 6. Pulmonary endothelial synthesis of vasoactive EET and HETE species. Further pathways of AA metabolism in the pulmonary endothelial cell can produce EET and HETE. The action of the cytochrome P450 epoxygenases 2C and 2J can produce a family of EET species, with various vasomotor and inflammatory functions, detailed in the text. The main catabolizer of EET species is the soluble epoxide hydrolase (sEH), which breaks EET species down into dihydroxyeicosatrienoic acids (DHET). Alternatively, ω or ω‐1 hydrolases can metabolize AA into 20‐ or 19‐HETE.


Figure 7. Pulmonary endothelial synthesis and secretion of ET‐1. ET‐1 is produced in the pulmonary endothelial cell in a prepro form, which is cleaved into pro‐ET‐1 (also called big ET‐1). The enzyme endothelin converting enzyme (ECE) then cleaves big ET‐1 into mature ET‐1, which then has both autocrine and paracrine functions within the lung vasculature. In the EC itself, ET‐1 can bind to the ETB receptor (ETBR), which results in nitric oxide (NO) and prostacyclin (PGI2) production. These two mediators can both subsequently act upon the smooth muscle cell to cause vasorelaxation. In a paracrine fashion, ET‐1 itself can bind to its receptors on smooth muscle cells, ETAR and ETBR, to cause G‐protein‐mediated activation of phospholipase C (PLC). The resultant production of inositol triphosphate (IP3) and diacylglycerol (DAG) results in SMC migration, and contraction, or proliferation, respectively.


Figure 8. Sources of reactive oxygen species in the pulmonary endothelial cell. A critically important source of ROS in the pulmonary EC is the family of NADPH oxidase (NOX) enzymes. Activated by angiotensin II, TGF‐β, and shear stress, NOX isoforms present in the lung can generate large quantities of ROS. Similarly, the mitochondrial electron transport chain can produce ROS, although controversy exists regarding whether hyperoxia or hypoxia favors this condition, and this controversy is discussed in the main text. Importantly, uncoupled eNOS produces ROS, and this ROS then leads to further eNOS uncoupling, which is an important propagator of inflammatory stress under a variety of pathological states. Elevated ROS levels leads to endothelial cytoskeletal changes, increased surface adhesion molecule expression, platelet adhesion, and a state referred to as endothelial dysfunction. Important enzymes such as superoxide dismutase exist to metabolize ROS and limit its accumulation.


Figure 9. Heterogeneous distribution of Weibel‐Pallade Bodies (WPB) in the lung endothelium. Representative electron micrographs of pulmonary endothelial cells from human lungs. WPB are indicated by arrowheads. Upper panels: High and low magnification images of extra‐alveolar endothelial cells, showing readily detectable WPB. Lower panels depict alveolar capillary endothelial cells, which are notably devoid of WPB. EC, endothelial cells, N, nucleus, BM, basement membrane, *, caveolae. Reproduced, with permission, from ().


Figure 10. Transcytosis of interleukin‐8 by endothelial cells. Stacked and reconstructed electron micrographs of human skin venular endothelial cells, with gold labeling of interleukin‐8 (IL‐8). (A) IL‐8 binding and uptake at the basolateral (abluminal) surface of the venular EC. (B) and (C) demonstrate IL‐8 in vesicles below the plasma membrane. (D) and (E) demonstrate IL‐8 now subsequently appearing at the luminal surface of the EC. (F) Colocalization of IL‐8 (10 nm gold) and caveolin‐1 (5 nm gold). Reproduced, with permission, from ().


Figure 11. Structure of the endothelial glycocalyx and endothelial surface layer (ESL). At the luminal surface of the pulmonary EC, the glycocalyx comprises a thin layer of plasma membrane glycoproteins and glycolipids. External to that is a thicker layer of glycosaminoglycans (GAG), heparan sulfate, hyaluronan, and other molecules, termed the ESL. These layers (A) separate the EC surface from the circulating plasma, (B) allow for mechanosensation and outside‐in signaling to the EC, (C) performs a critical role in maintaining the endothelial barrier, and (D) can be importantly degraded and modified during periods of inflammation.


Figure 12. The endothelial glycocalyx in vivo, and its degradation by LPS. (A) Schematic representation of the glycocalyx and endothelial surface layer (ESL). Note that the glycocalyx is not permeable to large molecules such as fluorescent dextrans. Therefore, measurement of vessel diameter by visible light microscopy versus fluorescence should allow for the measurement of the ESL. The images in the right of the panel show mouse sub‐pleural microvessels (MV) imaged with visible light (DIC) or fluorescence (FITC). The difference between the DIC and FITC width represents the ESL. (A) Alveolus. (B) Group data of calculated ESL thickness as in (A), taken from mice treated with LPS or TNF‐α, demonstrating how inflammatory stimuli can degrade the ESL. (C) These effects are not seen in mice lacking functional TNF receptors. Reproduced, with permission, from ().


Figure 13. Pulmonary endothelial angiotensin signaling. The majority of conversion of AngI to AngII is carried out by angiotensin converting enzyme (ACE), bound to the EC surface. AngII can then bind to AT1R or AT2R (left side). Signaling through AT1R results in G‐protein‐mediated release of IP3 and DAG, as well as stimulation of MAP kinases. The downstream effects include PASMC contraction and migration, as well as EC ROS production and vascular remodeling. Signaling via the AT2R is incompletely understood in the lung, but in general antagonizes the effects of AT1R signaling, leading to PASMC relaxation, and protection from adverse remodeling. The cell on the right depicts further metabolism of AngII by ACE2, producing Ang(1‐7), which subsequently binds the Mas receptor. Mas has a highly protective effect during states of acute lung injury and inflammation, decreasing ROS production, enhancing NO release, and protecting against thrombosis and EC apoptosis.


Figure 14. The conducted response in hypoxic pulmonary vasoconstriction. (A) Intravital microscopic images taken from wild‐type (top panels) or Cx40−/− mice (bottom panels) during normoxic or hypoxic ventilation, as indicated. Endothelial cells have been loaded with the voltage‐sensitive fluorophore, di‐8‐ANEPPS. Arteriolar vessel margins are indicated by dotted lines. Capillary endothelial cells are circled, while arteriolar EC are indicated by squares. Not that in wild‐type mice, depolarization in response to hypoxia occurs in both the capillary and arteriolar EC. However, in Cx40−/− mice, only the capillary EC depolarize, indicating the requirement of gap junction communication for the conducted response. (C) and (D) show group data describing the above. In (D), depolarization in response to hypoxia is not seen in the pulmonary arteriolar EC in Cx40−/− mice, indicative of the requirement for gap junctional communication. Reproduced, with permission, from ().


Figure 15. Schematic diagram representing the conducted response. It is hypothesized that hypoxia inhibits oxygen‐sensitive K+ channels, leading to EC depolarization at the alveolar capillary level. This depolarization spreads between EC in a retrograde fashion to the muscularized arteriolar EC via Cx40‐containing gap junctions. At this point, communication with the PASMC can occur, resulting in vasoconstriction. A contribution of myoendothelial gap junctions containing Cx43 and Cx40 is likely involved in this process.
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Neil M. Goldenberg, Wolfgang M. Kuebler. Endothelial Cell Regulation of Pulmonary Vascular Tone, Inflammation, and Coagulation. Compr Physiol 2015, 5: 531-559. doi: 10.1002/cphy.c140024