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Cytoskeletal Dynamics and Lung Fluid Balance

Stephen M. Vogel, Asrar B. Malik

10.1002/cphy.c100006

Source: Volume 2, Issue 1, January 2012

Published online: January 2012

Full Article on Wiley Online Library



Abstract

This article examines the role of the endothelial cytoskeleton in the lung's ability to restrict fluid and protein to vascular space at normal vascular pressures and thereby to protect lung alveoli from lethal flooding. The barrier properties of microvascular endothelium are dependent on endothelial cell contact with other vessel‐wall lining cells and with the underlying extracellular matrix (ECM). Focal adhesion complexes are essential for attachment of endothelium to ECM. In quiescent endothelial cells, the thick cortical actin rim helps determine cell shape and stabilize endothelial adherens junctions and focal adhesions through protein bridges to actin cytoskeleton. Permeability‐increasing agonists signal activation of “small GTPases” of the Rho family to reorganize the actin cytoskeleton, leading to endothelial cell shape change, disassembly of cortical actin rim, and redistribution of actin into cytoplasmic stress fibers. In association with calcium‐ and Src‐regulated myosin light chain kinase (MLCK), stress fibers become actinomyosin‐mediated contractile units. Permeability‐increasing agonists stimulate calcium entry and induce tyrosine phosphorylation of VE‐cadherin (vascular endothelial cadherin) and β‐catenins to weaken or pull apart endothelial adherens junctions. Some permeability agonists cause latent activation of the small GTPases, Cdc42 and Rac1, which facilitate endothelial barrier recovery and eliminate interendothelial gaps. Under the influence of Cdc42 and Rac1, filopodia and lamellipodia are generated by rearrangements of actin cytoskeleton. These motile evaginations extend endothelial cell borders across interendothelial gaps, and may initiate reannealing of endothelial junctions. Endogenous barrier protective substances, such as sphingosine‐1‐phosphate, play an important role in maintaining a restrictive endothelial barrier and counteracting the effects of permeability‐increasing agonists. © 2012 American Physiological Society. Compr Physiol 2:449‐478, 2012.

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Figure 1. Figure 1.

Transition between quiescent and active endothelial phenotypes. Left: Endothelial cell activation by permeability‐increasing agonist. Binding of thrombin to its receptor increases intracellular Ca2+, which forms a complex with calmodulin (CaM) that can activate myosin light chain kinase (MLCK). Phosphorylated form of myosin light chain (MLC) stimulates actinomyosin‐based endothelial contractility, which leads to the formation of interendothelial gaps. Examples of regulatory molecules that increase or decrease endothelial contractility are depicted in green or white, respectively: Src kinases (pp60src) increase MLCK activity by tyrosine phosphorylation; rho kinase augments EC contractility by inhibiting both MLC phosphatase and the actin‐depolymerizing protein, cofilin; p38 mitogen‐activated protein kinase (MAPK) inhibits heat shock protein (HSP) 27 activity to promote actin stress fiber formation, as does the actin‐associated capping/severing protein gelsolin. Right: Quiescent endothelial cells possess a thick cortical actin rim, and adhere tightly to each other via interendothelial junctional complexes (tight junctions and adherens junctions) and to the underlying ECM via focal adhesions. Tight junctions consist of transmembrane occludin proteins linked to the endothelial actin cytoskeleton by proteins of the zona occludins family (ZO‐1). Adherens junctions mediate cell‐cell contact through homotypic binding between the extracellular domains of endothelial‐specific VE‐cadherin, and are stabilized by protein bridges between the cytoplasmic tail of VE‐cadherin and cortical actin rim that include, among other components, β/γ‐catenins (plakoglobin) and α‐catenin. The tyrosine phosphatase SHP2 may contribute to stability of adherens junctions by decreasing tyrosine phosphorylation of catenins. Platelet endothelial cell adhesion molecule‐1 (PECAM‐1) is concentrated in intercellular clefts, but it is not associated specifically with adherens or tight junctions; PECAM‐1 can mediate homotypic or heterotypic binding to an adjacent cell, and is internally tethered to cortical actin rim 17,187. Focal adhesion plaques consisting of talin, paxillin (Pax), vinculin (Vin), α‐actinin (A), and focal adhesion kinase (FAK) link endothelial integrins to cortical actin rim to stabilize focal adhesions. Sphingosine‐1‐phosphate (S‐1‐P) binding to its receptor promotes quiescent endothelial phenotype and counteracts the effects of permeability‐increasing agonists such as thrombin. PAR‐1, protein‐activated receptor‐1; EDG receptor, endothelial differentiation gene receptor. American Physiological Society, used with permission 61.
Figure 2. Figure 2.

Separate fluid and protein transport pathways through microvessel endothelium. Fluid moves through paracellular pathway according to Starling forces. Protein is transported through endothelial cells via vesicles or transendothelial channels. Because the protein and fluid pathways are separated, local protein gradients can theoretically develop in the interstitium. American Physiological Society, used with permission 174.
Figure 3. Figure 3.

The vesicular protein‐transport pathway in pulmonary microvascular endothelium. Main electron micrograph shows labeling of endothelial vesicles with the vascular tracer dinitrophenol‐conjugated albumin (A‐DNP) in a murine postcapillary venule. Inset image is similar except that it depicts albumin transport in a murine lung capillary. Albumin detection was with an electron opaque anti‐DNP antibody. Note abundant labeling of endothelial caveolae in both microvessels at various stages of albumin transcytosis and absence of the tracer molecule (diameter 12 nm) from interendothelial space (see inset). Calibration bars are 100 nm (main image) and 80 nm (inset). PVS, perivascular space. American Physiological Society, used with permission 217.
Figure 4. Figure 4.

Starling forces acting on lung microvessel walls. P, π, and σ denote hydrostatic pressure, oncotic pressure, and osmotic reflection coefficient, respectively. ΔP is the net pulmonary driving pressure, which is normally outward. In mild hydrostatic edema, capillary pressure is elevated, leading to an intertstitial edema. In hypoproteinemia, oncotic pressure difference is reduced causing an interstitial edema. Int, interstitium; cap, capillary.
Figure 5. Figure 5.

Formation of protein‐rich pulmonary edema fluid. Normal and injured alveolar‐capillary barriers are illustrated in left and right panels. In the acute phase of ARDS (acute respiratory distress syndrome), capillary endothelium is damaged and neutrophils adherent to injured capillary endothelium migrate through vessel wall and into alveolar space. Endothelial swelling and blebbing takes place and prominent interendothelial gaps form, thereby compromising the endothelial barrier to blood cells and plasma. Platelets aggregate on damaged vessel wall. Pulmonary interstitium is dilated due to fluid accumulation. Alveolar macrophages release interleukins (IL‐1, 6, 8, and 10) and TNFα, further stimulating neutrophil margination and migration. IL‐1 can also stimulate interstitial fibroblasts to generate ECM proteins. Neutrophils release oxidants, proteases, leukotrienes, and platelet‐activating factor. Damaged epithelial cells become detached from basement membrane and are replaced with proteinaceous hyaline “membranes.” As the alveolar‐capillary barrier deteriorates, increasing numbers of alveoli become flooded with fluid rich in plasma proteins, which inactivates surfactant. Hence, loss of alveolar‐epithelial barrier function leads to protein‐rich edema. MIF, macrophage inhibitory factor. Reprinted with permission of the American Thoracic Society. Copyright© American Thoracic Society 160.
Figure 6. Figure 6.

Actin microfilaments. (A) Electron micrograph of an actin microfilament decorated with myosin heads. (B) Model of elongation of microfilament. Ratio of dissociation rate constant (s−1) to association rate constant (μM−1 s−1) gives K, the equilibrium dissociation constant (μM). ATP‐bound actin monomer has a higher affinity for barbed end than for pointed end. Reprinted, with permission, from reference 212.
Figure 7. Figure 7.

Organization of microtubule cytoskeleton in monolayer of human lung microvascular endothelial cells (HLMVECs). (A) Microtubules (yellow) extend from the microtubule‐organizing center (MTOC) to cell periphery; nuclei are stained in blue (Image courtesy of Dr. Yulia Komarova). (B) Schematic diagram of the microtubule (MT) cytoskeleton. Minus ends of MTs attach to centrosome (yellow) and plus ends grow radially toward cell periphery. MTs serve as tracks for mobile vesicles, which move from centrosome to cell periphery (red circles) or backwards toward the centrosome (green circles). Reprinted, with permission, from reference 108.
Figure 8. Figure 8.

Growth and shrinkage of microtubules (MT). Presence of GTP–(α, β)‐tubulin (pink) at the plus end stabilizes the growth phase of MT. GTP hydrolysis results in rapid MT shrinkage, with individual protofilaments bending away from the microtubule axis. The transition from growth to shrinkage is referred to as catastrophe; transition from shrinkage to growth is known as rescue. Reprinted, with permission, from reference 109.
Figure 9. Figure 9.

Activation of LIM kinase 1 (LIMK1) destabilizes pulmonary interendothelial junctions. (A and B) Electron micrographs of interendothelial junctions (arrows) in WT (A) and limk1–/– mice (B) without and with PAR‐1 peptide agonist as indicated. Magnification, 60,000. (C) Morphometric analysis of open interendothelial junctions induced by exposure of mouse lungs to PAR‐1 agonist. Each bar shows percentage of open junctions out of 50 observations made. PAR‐1 agonist induced significantly more open junctions in WT lungs (P = 0.010197). Error bars represent SEM. Reprinted, with permission, from reference 92.
Figure 10. Figure 10.

Interendothelial junctions in quiescent cells. (A) Diagram of endothelial adherens junctions and their relationship to actin cytoskeleton. Homotypic adhesion between extracellular domains (EXDS) of VE‐cadherin molecules joins adjacent cells. The juxtramembrane domain (JMD) of VE‐cadherin binds p120 (aqua), a protein regulating the stability of adherens junctions. The carboxyl‐terminal domain (CTD) binds exclusively to either β‐catenin or γ‐catenin (plakoglobin; green), which in turn binds to α‐catenin (dark blue). α‐Catenin interacts with F‐actin directly and indirectly through binding to α‐actinin (light orange) to create a strong link between VE‐cadherin and actin cytoskeleton. Vinculin, another actin‐associated protein, is also depicted (violet). (B) Diagram illustrating endothelial tight junctions. Homotypic adhesion between EXDs of occludin molecules (green) creates the tight junctional contact. The cytoplasmic carboxyl domain of occludin binds to ZO‐1 (red), which is indirectly linked to actin cytoskeleton via α‐catenin (dark blue). Reprinted, with permission, from reference 213.
Figure 11. Figure 11.

Organization of actin and microtubule cytoskeletons and their relationship to adherens junctions. Monolayer of quiescent human lung microvascular endothelial cells was stained for VE‐cadherin (red), α‐tubulin and actin (green, as indicated), and DNA (blue). Actin organizes into peripheral actin bundles along the VE‐cadherin‐mediated adhesions. Microtubules extend from the centrosome throughout the cell. Image courtesy of Dr.Yulia Komarova.
Figure 12. Figure 12.

Organization of focal adhesions. (A) Immunostaining of endothelial monolayer for actin (green) and focal‐adhesion protein, vinculin (red). (B) Diagram of the focal adhesion complex, showing integrin bound to ECM protein and its intracellular linkage to F‐actin. Image courtesy of Dr. Richard Minshall.
Figure 13. Figure 13.

Receptor control of microvessel permeability. Permeability‐increasing agonist such as thrombin, ligates proteinase‐activated receptor‐1 (PAR‐1), stimulating Gαq and Gα12/13. Gαq triggers increased intracellular Ca2+, which activates EC MLCK (endothelial‐cell myosin light chain kinase) and hence elevates the phosphorylation level of MLC (MLC‐P). Increased intracellular Ca2+ also activates protein kinases, such as PKCα (not shown), which phosphorylate GDI‐1 (GDP‐dissociation inhibitor) and p115RhoGEF (guanine nucleotide exchange factor), thereby increasing the activity of RhoA GTPase. RhoA activates ROCK (rho kinase) and LIMK1 (LIM kinase‐1), which regulates actin binding proteins (cofilin) and thus the state of actin polymerization. ROCK by phosphorylating MLC phosphatase (Ppase) inhibits Ppase and thus increases MLC‐P. Together, these events lead to increased actin‐myosin cross‐bridging, resulting in increased endothelial contraction. Image courtesy of Dr. Dolly Mehta.
Figure 14. Figure 14.

Mechanism of lamellipodia and filopodia formation. (A) Organization of actin cytoskeleton in lamellipodia of fish epidermal keratocytes. Reproduced, with permission, from reference 235. (B and C) Organization of actin cytoskeleton in filopodia embedded in lamellipodium in B16F1 melanoma cell. (B) Fluorescent image. Region enclosed by rectangle is enlarged in insets. Reproduced, with permission, from reference 171. (C) Platinum replica EM. Filopodium contains a tight bundle of actin filaments. Inset shows branched actin on surface of lamellipodium. Reproduced, with permission, from reference 261. (D and E) Model for mechanism of lamellipodial (top) and filopodial (bottom) protrusion depending on capping activity in cells. Top: When capping activity dominates either by activation of CP (capping protein) or inhibition of capping antagonists such as Ena/VASP‐like protein (enabled/ vasodilator‐stimulated phosphoprotein), filaments elongate for a brief period before becoming capped and, as a consequence, these filaments are relatively short. To maintain protrusion, capped filaments are replaced by newly nucleated side branches produced by Arp2/3 complex, favoring the formation of a branching, lamellipodial network. Bottom: Low‐capping activity resulting either from inhibition of CP or activation of proteins that promote anticapping and antibranching such as Ena/VASP, favor filament elongation leading to long and unbranched filaments, which converge and subsequently become bundled by actin cross‐linking molecule, fascin. Filopodia formation is therefore predominant. Modified, with permission, from reference 171.
Figure 15. Figure 15.

Thrombin‐induced stress fiber formation in endothelial cells (murine lung). Actin is stained in green and nuclei, in blue. Image courtesy of Dr. C. Tiruppathi.
Figure 16. Figure 16.

Intracellular distribution of VE‐cadherin in resting endothelial cells and in cells challenged with pro‐inflammatory mediator, thrombin. Confluent monolayers of HLMVECs, either untreated or treated with 50 nM human α‐thrombin for 30 min, were stained for VE‐cadherin (yellow) and DNA (blue). Thrombin induces disruption of VE‐cadherin‐mediated adhesions and formation of gaps between cells. (Images courtesy of Emily Vandenbroucke).


Figure 1.

Transition between quiescent and active endothelial phenotypes. Left: Endothelial cell activation by permeability‐increasing agonist. Binding of thrombin to its receptor increases intracellular Ca2+, which forms a complex with calmodulin (CaM) that can activate myosin light chain kinase (MLCK). Phosphorylated form of myosin light chain (MLC) stimulates actinomyosin‐based endothelial contractility, which leads to the formation of interendothelial gaps. Examples of regulatory molecules that increase or decrease endothelial contractility are depicted in green or white, respectively: Src kinases (pp60src) increase MLCK activity by tyrosine phosphorylation; rho kinase augments EC contractility by inhibiting both MLC phosphatase and the actin‐depolymerizing protein, cofilin; p38 mitogen‐activated protein kinase (MAPK) inhibits heat shock protein (HSP) 27 activity to promote actin stress fiber formation, as does the actin‐associated capping/severing protein gelsolin. Right: Quiescent endothelial cells possess a thick cortical actin rim, and adhere tightly to each other via interendothelial junctional complexes (tight junctions and adherens junctions) and to the underlying ECM via focal adhesions. Tight junctions consist of transmembrane occludin proteins linked to the endothelial actin cytoskeleton by proteins of the zona occludins family (ZO‐1). Adherens junctions mediate cell‐cell contact through homotypic binding between the extracellular domains of endothelial‐specific VE‐cadherin, and are stabilized by protein bridges between the cytoplasmic tail of VE‐cadherin and cortical actin rim that include, among other components, β/γ‐catenins (plakoglobin) and α‐catenin. The tyrosine phosphatase SHP2 may contribute to stability of adherens junctions by decreasing tyrosine phosphorylation of catenins. Platelet endothelial cell adhesion molecule‐1 (PECAM‐1) is concentrated in intercellular clefts, but it is not associated specifically with adherens or tight junctions; PECAM‐1 can mediate homotypic or heterotypic binding to an adjacent cell, and is internally tethered to cortical actin rim 17,187. Focal adhesion plaques consisting of talin, paxillin (Pax), vinculin (Vin), α‐actinin (A), and focal adhesion kinase (FAK) link endothelial integrins to cortical actin rim to stabilize focal adhesions. Sphingosine‐1‐phosphate (S‐1‐P) binding to its receptor promotes quiescent endothelial phenotype and counteracts the effects of permeability‐increasing agonists such as thrombin. PAR‐1, protein‐activated receptor‐1; EDG receptor, endothelial differentiation gene receptor. American Physiological Society, used with permission 61.



Figure 2.

Separate fluid and protein transport pathways through microvessel endothelium. Fluid moves through paracellular pathway according to Starling forces. Protein is transported through endothelial cells via vesicles or transendothelial channels. Because the protein and fluid pathways are separated, local protein gradients can theoretically develop in the interstitium. American Physiological Society, used with permission 174.



Figure 3.

The vesicular protein‐transport pathway in pulmonary microvascular endothelium. Main electron micrograph shows labeling of endothelial vesicles with the vascular tracer dinitrophenol‐conjugated albumin (A‐DNP) in a murine postcapillary venule. Inset image is similar except that it depicts albumin transport in a murine lung capillary. Albumin detection was with an electron opaque anti‐DNP antibody. Note abundant labeling of endothelial caveolae in both microvessels at various stages of albumin transcytosis and absence of the tracer molecule (diameter 12 nm) from interendothelial space (see inset). Calibration bars are 100 nm (main image) and 80 nm (inset). PVS, perivascular space. American Physiological Society, used with permission 217.



Figure 4.

Starling forces acting on lung microvessel walls. P, π, and σ denote hydrostatic pressure, oncotic pressure, and osmotic reflection coefficient, respectively. ΔP is the net pulmonary driving pressure, which is normally outward. In mild hydrostatic edema, capillary pressure is elevated, leading to an intertstitial edema. In hypoproteinemia, oncotic pressure difference is reduced causing an interstitial edema. Int, interstitium; cap, capillary.



Figure 5.

Formation of protein‐rich pulmonary edema fluid. Normal and injured alveolar‐capillary barriers are illustrated in left and right panels. In the acute phase of ARDS (acute respiratory distress syndrome), capillary endothelium is damaged and neutrophils adherent to injured capillary endothelium migrate through vessel wall and into alveolar space. Endothelial swelling and blebbing takes place and prominent interendothelial gaps form, thereby compromising the endothelial barrier to blood cells and plasma. Platelets aggregate on damaged vessel wall. Pulmonary interstitium is dilated due to fluid accumulation. Alveolar macrophages release interleukins (IL‐1, 6, 8, and 10) and TNFα, further stimulating neutrophil margination and migration. IL‐1 can also stimulate interstitial fibroblasts to generate ECM proteins. Neutrophils release oxidants, proteases, leukotrienes, and platelet‐activating factor. Damaged epithelial cells become detached from basement membrane and are replaced with proteinaceous hyaline “membranes.” As the alveolar‐capillary barrier deteriorates, increasing numbers of alveoli become flooded with fluid rich in plasma proteins, which inactivates surfactant. Hence, loss of alveolar‐epithelial barrier function leads to protein‐rich edema. MIF, macrophage inhibitory factor. Reprinted with permission of the American Thoracic Society. Copyright© American Thoracic Society 160.



Figure 6.

Actin microfilaments. (A) Electron micrograph of an actin microfilament decorated with myosin heads. (B) Model of elongation of microfilament. Ratio of dissociation rate constant (s−1) to association rate constant (μM−1 s−1) gives K, the equilibrium dissociation constant (μM). ATP‐bound actin monomer has a higher affinity for barbed end than for pointed end. Reprinted, with permission, from reference 212.



Figure 7.

Organization of microtubule cytoskeleton in monolayer of human lung microvascular endothelial cells (HLMVECs). (A) Microtubules (yellow) extend from the microtubule‐organizing center (MTOC) to cell periphery; nuclei are stained in blue (Image courtesy of Dr. Yulia Komarova). (B) Schematic diagram of the microtubule (MT) cytoskeleton. Minus ends of MTs attach to centrosome (yellow) and plus ends grow radially toward cell periphery. MTs serve as tracks for mobile vesicles, which move from centrosome to cell periphery (red circles) or backwards toward the centrosome (green circles). Reprinted, with permission, from reference 108.



Figure 8.

Growth and shrinkage of microtubules (MT). Presence of GTP–(α, β)‐tubulin (pink) at the plus end stabilizes the growth phase of MT. GTP hydrolysis results in rapid MT shrinkage, with individual protofilaments bending away from the microtubule axis. The transition from growth to shrinkage is referred to as catastrophe; transition from shrinkage to growth is known as rescue. Reprinted, with permission, from reference 109.



Figure 9.

Activation of LIM kinase 1 (LIMK1) destabilizes pulmonary interendothelial junctions. (A and B) Electron micrographs of interendothelial junctions (arrows) in WT (A) and limk1–/– mice (B) without and with PAR‐1 peptide agonist as indicated. Magnification, 60,000. (C) Morphometric analysis of open interendothelial junctions induced by exposure of mouse lungs to PAR‐1 agonist. Each bar shows percentage of open junctions out of 50 observations made. PAR‐1 agonist induced significantly more open junctions in WT lungs (P = 0.010197). Error bars represent SEM. Reprinted, with permission, from reference 92.



Figure 10.

Interendothelial junctions in quiescent cells. (A) Diagram of endothelial adherens junctions and their relationship to actin cytoskeleton. Homotypic adhesion between extracellular domains (EXDS) of VE‐cadherin molecules joins adjacent cells. The juxtramembrane domain (JMD) of VE‐cadherin binds p120 (aqua), a protein regulating the stability of adherens junctions. The carboxyl‐terminal domain (CTD) binds exclusively to either β‐catenin or γ‐catenin (plakoglobin; green), which in turn binds to α‐catenin (dark blue). α‐Catenin interacts with F‐actin directly and indirectly through binding to α‐actinin (light orange) to create a strong link between VE‐cadherin and actin cytoskeleton. Vinculin, another actin‐associated protein, is also depicted (violet). (B) Diagram illustrating endothelial tight junctions. Homotypic adhesion between EXDs of occludin molecules (green) creates the tight junctional contact. The cytoplasmic carboxyl domain of occludin binds to ZO‐1 (red), which is indirectly linked to actin cytoskeleton via α‐catenin (dark blue). Reprinted, with permission, from reference 213.



Figure 11.

Organization of actin and microtubule cytoskeletons and their relationship to adherens junctions. Monolayer of quiescent human lung microvascular endothelial cells was stained for VE‐cadherin (red), α‐tubulin and actin (green, as indicated), and DNA (blue). Actin organizes into peripheral actin bundles along the VE‐cadherin‐mediated adhesions. Microtubules extend from the centrosome throughout the cell. Image courtesy of Dr.Yulia Komarova.



Figure 12.

Organization of focal adhesions. (A) Immunostaining of endothelial monolayer for actin (green) and focal‐adhesion protein, vinculin (red). (B) Diagram of the focal adhesion complex, showing integrin bound to ECM protein and its intracellular linkage to F‐actin. Image courtesy of Dr. Richard Minshall.



Figure 13.

Receptor control of microvessel permeability. Permeability‐increasing agonist such as thrombin, ligates proteinase‐activated receptor‐1 (PAR‐1), stimulating Gαq and Gα12/13. Gαq triggers increased intracellular Ca2+, which activates EC MLCK (endothelial‐cell myosin light chain kinase) and hence elevates the phosphorylation level of MLC (MLC‐P). Increased intracellular Ca2+ also activates protein kinases, such as PKCα (not shown), which phosphorylate GDI‐1 (GDP‐dissociation inhibitor) and p115RhoGEF (guanine nucleotide exchange factor), thereby increasing the activity of RhoA GTPase. RhoA activates ROCK (rho kinase) and LIMK1 (LIM kinase‐1), which regulates actin binding proteins (cofilin) and thus the state of actin polymerization. ROCK by phosphorylating MLC phosphatase (Ppase) inhibits Ppase and thus increases MLC‐P. Together, these events lead to increased actin‐myosin cross‐bridging, resulting in increased endothelial contraction. Image courtesy of Dr. Dolly Mehta.



Figure 14.

Mechanism of lamellipodia and filopodia formation. (A) Organization of actin cytoskeleton in lamellipodia of fish epidermal keratocytes. Reproduced, with permission, from reference 235. (B and C) Organization of actin cytoskeleton in filopodia embedded in lamellipodium in B16F1 melanoma cell. (B) Fluorescent image. Region enclosed by rectangle is enlarged in insets. Reproduced, with permission, from reference 171. (C) Platinum replica EM. Filopodium contains a tight bundle of actin filaments. Inset shows branched actin on surface of lamellipodium. Reproduced, with permission, from reference 261. (D and E) Model for mechanism of lamellipodial (top) and filopodial (bottom) protrusion depending on capping activity in cells. Top: When capping activity dominates either by activation of CP (capping protein) or inhibition of capping antagonists such as Ena/VASP‐like protein (enabled/ vasodilator‐stimulated phosphoprotein), filaments elongate for a brief period before becoming capped and, as a consequence, these filaments are relatively short. To maintain protrusion, capped filaments are replaced by newly nucleated side branches produced by Arp2/3 complex, favoring the formation of a branching, lamellipodial network. Bottom: Low‐capping activity resulting either from inhibition of CP or activation of proteins that promote anticapping and antibranching such as Ena/VASP, favor filament elongation leading to long and unbranched filaments, which converge and subsequently become bundled by actin cross‐linking molecule, fascin. Filopodia formation is therefore predominant. Modified, with permission, from reference 171.



Figure 15.

Thrombin‐induced stress fiber formation in endothelial cells (murine lung). Actin is stained in green and nuclei, in blue. Image courtesy of Dr. C. Tiruppathi.



Figure 16.

Intracellular distribution of VE‐cadherin in resting endothelial cells and in cells challenged with pro‐inflammatory mediator, thrombin. Confluent monolayers of HLMVECs, either untreated or treated with 50 nM human α‐thrombin for 30 min, were stained for VE‐cadherin (yellow) and DNA (blue). Thrombin induces disruption of VE‐cadherin‐mediated adhesions and formation of gaps between cells. (Images courtesy of Emily Vandenbroucke).

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Further Reading
 1. Prasain N, Stevens T. The actin cytoskeleton in endothelial cell phenotypes. Microvasc Res 77: 53‐63, 2009.
 2. Romer LH, Birukov KG, Garcia JG. Focal adhesions: Paradigm for a signaling nexus. Circ Res 98: 606‐616, 2006.
 3. Wang L, Dudek SM. Regulation of vascular permeability by sphingosine 1‐phosphate Microvasc Res 77: 39‐45, 2009.
 4. Dejana E, Orsenigo F, Lampugnani MG. The role of adherens junctions and VE‐cadherin in the control of vascular permeability. J Cell Sci 121(Pt 13): 2115‐2122, 2008.
 5. Mehta D, Malik AB. Signaling mechanisms regulating endothelial permeability. Physiol Rev 86: 279‐367, 2006.
 6. Bazzoni G, Dejana E. Endothelial cell‐to‐cell junctions: molecular organization and role in vascular homeostasis. Physiol Rev 84: 869‐901, 2004.
 7. Csortos C, Kolosova I, Verin AD. Regulation of vascular endothelial cell barrier function and cytoskeleton structure by protein phosphatases of the PPP family. Am J Physiol Lung Cell Mol Physiol 293: L843‐L854, 2007.
 8. Komarova YA, Mehta D, Malik AB. Dual regulation of endothelial junctional permeability. Sci STKE. 2007: re8, 2007.

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Article Title: Cytoskeletal Dynamics and Lung Fluid Balance
 

How to Cite

Stephen M. Vogel, Asrar B. Malik. Cytoskeletal Dynamics and Lung Fluid Balance. Compr Physiol 2012, 2: 449-478. doi: 10.1002/cphy.c100006