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Microcirculatory Exchange Function

Walter N. Durán, Fabiola A. Sánchez, Jerome W. Breslin

10.1002/cphy.cp020404

Source: Supplement 9: Handbook of Physiology, The Cardiovascular System, Microcirculation

Originally published: 2008

Published online: January 2011

Full Article on Wiley Online Library



Abstract

The sections in this article are:

1 Introduction
2 Physical Principles Controlling Fluid and Solute Transport
2.1 General Characteristics of Fluid and Solute Flux
2.2 Exchange Microvessels
2.3 Experimental Models for Studying Microvascular Transport
2.4 Paracellular and Transcellular Transport
3 Regulatory Mechanisms
3.1 Agonists and Receptors: Localization and Function
3.2 Signal Transduction Pathways
4 Concluding Remarks
Figure 1. Figure 1.

Microvascular hyperpermeability response to platelet activating factor (PAF: 1.0 × 10−7 M) in the hamster cheek pouch. The details of the microvascular network are observed by fluorescence intravital microscopy using a macromolecular fluorochrome (i.e. FITC‐dextran 150kDa). The branching pattern of the arterioles (a) and tributaries to venules (v) can be followed easily. The control panel (A) shows absence of FITC‐dextran 150 in the interstitium. Topical application of PAF (B) induced arteriolar constriction and permeation of the fluorescent macromolecular tracer into the interstitium across postcapillary venules (arrows). Modified from 66 with permission. (See page 3 in colour section at the back of the book)
Figure 2. Figure 2.

The Landis technique. A single microvessel is occluded with a glass micropipette and the distance between an erythrocyte trapped and in the microvessel lumen and the occluding probe is measured as a function of time. The microvessel diameter is also measured. Volume flux (J,JS) is estimated using the initial erythrocyte velocity. Hydraulic conductivity (Lp) is determined as the slope of J,JS as a function of mean occluded intraluminal pressure (P).
Figure 3. Figure 3.

The isolated venule technique. (A) Schematic of the experimental setup. A venule is dissected, excised, and mounted on two glass micropipettes and bathed with an albumin physiological saline solution at 37°C. Pressure and flow are controlled by altering the heights of the inflow and outflow reservoirs, which are mounted on manometers. (B) To assess permeability, the fluorescence intensity of a window containing the venule and the nearby extraluminal area is measured. The inflow is quickly switched from normal APSS to APSS containing a fluorochrome‐labeled tracer (e.g. FITC‐albumin), causing a step increase fluorescence intensity (ΔIfo) that is proportional to the number of solute tracer molecules that have entered the lumen. This is followed by a gradual increase in intensity (dIf/dt)o. The rate of (dIf/dt)o is proportional to solute transport across the microvascular wall. The intensity returns to the basal level when the tracer is washed out from the lumen.

Adapted from 34 and 79 with permission
Figure 4. Figure 4.

Cultured endothelial cell (EC) permeability models. An EC monolayer is grown on a porous membrane that is placed between two chambers. (A) A vertical chamber system is shown, in which the upper chamber represents the “luminal” side and the tower chamber represents the “abluminal” side of the endothelial monolayer. Note that the fluid levels of the upper and lower chambers are kept at the same height to minimize any hydrostatic differences between the two chambers. (B) A horizontal setup.
Figure 5. Figure 5.

Ultrastructure of canine cardiac postcapillary venule. The electron micrograph demonstrates the complex morphology of endothelial cells (ECs). Possible transport pathways for solutes are indicated by different arrows. Caveolae and vesicles are abundant throughout the EC. WP. Weibel—Palade body. Durán, WN and Berendsen P (unpublished). (See page 3 in colour section at the back of the book)
Figure 6. Figure 6.

Routes of fluid and solute transport across the endothelium. Two pathways, one paracellular and one transcellular, have been proposed. In the paracellular pathway, fluid and solutes pass between endothelial cells (ECs), with the major barrier being the tightness of junctions between ECs. Several molecules thought to regulate junctional integrity are shown. In the transcellular pathway, active vesicular transport, or the formation of channels of VVOs, is proposed as a route for fluid and solute passage.
Figure 7. Figure 7.

Signal transduction pathways in microvascular permeability. Receptors are activated upon binding to their respective agonists and interact with downstream signaling pathways. The diagram reflects a typical set of cascades activated by permeability agonists; VEGF is used as an example.
Figure 8. Figure 8.

Transfection of ca‐ROCK protein increases coronary venular permeability. Isolated porcine coronary venules were transfected with 3 μg/ml ca‐ROCK protein for 120min in absence (solid line, squares, n = 6) or presence (dashed line, triangles, n = 4) of the ROCK inhibitor Y‐27632 (5 × 10−6M). Y‐27632 was added to the bath 20min prior to the addition of ca‐ROCK. Transfection was initiated at 0min and continued throughout the time course. *Indicates P < 0.05 vs. baseline permeability (0‐min time point). ‡Indicates P < 0.05 vs. ca‐ROCK transfection when Y‐27632 is present. From Ref. 307 with permission.
Figure 9. Figure 9.

MLC phosphorylation as a mechanism of endothelial barrier regulation. Inflammatory stimuli cause elevated phosphorylation of MLC on Thr‐18/Ser‐19 in endothelial cells, which in turn elevates actin—myosin mediated contraction. This leads to increased cellular centrifugal force that promotes opening of endothelial cell—cell junctions, causing increased endothelial permeability. MLC phosphorylation increases when MLCK activity is elevated or MLCP activity is reduced. The Rho/ROCK pathway can inactivate MLCP through phosphorylation of the MLCP targeting subunit MYPT‐1 on Thr‐696. (See page 4 in colour section at the back of the book)
Figure 10. Figure 10.

ROCK‐mediated tension and barrier dysfunction in coronary venular endothelial cell (EC) monolayers. Transfection of ca‐ROCK (3 μg/ml) into cultured coronary venular EC (CVEC) monolayers increases isometric tension (triangles) and causes a drop in TER (solid line). A time course overlay of ca‐ROCK‐stimulated changes in TER and tension shows that these two events occur simultaneously. From Ref. 307 with permission.
Figure 11. Figure 11.

Adherens junctions in endothelial cells (EC). (A) Adhesion between ECs is achieved by homotypic binding of VE‐cadherin molecules at endothelial cell—cell junctions. Several catenins (p120, α, β, γ) are associated with VE‐cadherin and regulate its homotypic binding, as well as linkage to the cytoskeleton. (B) Under stimulated conditions, such as during VEGF treatment, VE‐cadherin, β‐catenin, and p120 catenin are phosphorylated on tyrosine residues, and a reorganization of endothelial cell—cell junctions occurs, characterized by the appearance of several VE‐cadherin rich finger‐like projections (arrows). Scale bar represents 20μm. Images from Ref. 207 with permission. (See page 4 in colour section at the back of the book)
Figure 12. Figure 12.

PAF‐induced NO production. (A) NO concentration was measured in the suffusate of the hamster cheek pouch using a Nitric Oxide Analyzer 407. (B) Perivenular NO concentration was determined using a NO‐sensitive electrode in the mouse cremaster muscle. NO concentration rose quickly upon topical administration of PAF in both tissues. “From 407 with permission.
Figure 13. Figure 13.

Obligatory role of eNOS‐mediated NO production in PAF‐induced hyperpermeability in vivo. PAF‐induced hyperpermeability is markedly reduced in eNOS−/−mice compared to wild‐type control mice. Typical intravital fluorescence microscopy images of the respective cremaster muscles are shown below the IOI graph.

Adapted from Refs 74 and 410 with permission. (See page 4 in colour section at the back of the book)
Figure 14. Figure 14.

Translocation of eNOS. Top Panel: (A) Isolation of lipid rafts domains was done in control, Ach, or PAF‐treated cells. Fractions were probed against caveolin‐1 and eNOS. (A) ACh treatment. (B) PAF treatment. The blots represent three independent experiments. Bottom Panel: The influence of PAF and ACh on the distribution of eNOS‐GFP was assessed by fluorescence of GFP. The images are representative of three independent experiments. (A) ECV304‐eNOS‐GFP. (B) Human dermal microvascular cells stained with anti‐eNOS (red) and anti‐caveolin‐1 (green) antibodies. From Ref. 434 with permission. (See page 5 in colour section at the back of the book)
Figure 15. Figure 15.

Diagram of eNOS translocation process and eNOS activation. (A) Shows a simplified diagram of the eNOS translocation hypothesis. eNOS is anchored in caveolae through myristoylation and palmitoylation, and is associated with caveolin‐1. Upon stimulation with agonists, eNOS becomes phosphorylated, dissociates from caveolin‐1, and translocates from the caveolae to either the Golgi apparatus or the cytosol. It is postulated that preferential translocation to different compartments is related to vascular function and may allow eNOS to interact with specific function‐related target molecules. Even though the drawing depicts eNOS as being “free” in the cytosol, our recent‐unpublished data indicate that eNOS may be internalized to its intracellular targets via caveolae. (B). Several signals can regulate eNOS activity. While association with caveolin‐1 and phosphorylation of Thr‐497 decrease enzyme activity (NO production), interaction with Ca2+‐calmodulin and HSP90, phosphorylation on Ser‐1177 by Akt, and PKC activation can all increase activity. Panel A redrawn from 410 with permission.
Figure 16. Figure 16.

PAF‐induced eNOS phosphorylation and hyperpermeability in ECV304‐eNOSGFP cells. (A) ACh and PAF activate eNOS by Ser‐1177 phosphorylation and Thr495 dephosphorylation. (B) Inhibition of muscarinic ACh and PAF receptors blocks Ser‐1177 phosphorylation β‐actin served as a (loading) control in experiments shown in (A) and (B). (C) PAF increases permeability in ECVeNOS‐GFP cells. Monolayers of ECVeNOS‐GFP cells were treated with PAF and permeability to FITC‐Dx‐70 was measured. Data are expressed as permeability coefficients (mean ± SEM) for control and PAF‐stimulated monolayers. *P < 0.05, n = 6. From 434 with permission.
Figure 17. Figure 17.

Functions of Rap‐1. The diagram shows known and putative effectors and targets of Rap‐1. Arrows indicate stimulation, blunt ends denote inhibition.

Adapted from Ref. 457 with permission


Figure 1.

Microvascular hyperpermeability response to platelet activating factor (PAF: 1.0 × 10−7 M) in the hamster cheek pouch. The details of the microvascular network are observed by fluorescence intravital microscopy using a macromolecular fluorochrome (i.e. FITC‐dextran 150kDa). The branching pattern of the arterioles (a) and tributaries to venules (v) can be followed easily. The control panel (A) shows absence of FITC‐dextran 150 in the interstitium. Topical application of PAF (B) induced arteriolar constriction and permeation of the fluorescent macromolecular tracer into the interstitium across postcapillary venules (arrows). Modified from 66 with permission. (See page 3 in colour section at the back of the book)



Figure 2.

The Landis technique. A single microvessel is occluded with a glass micropipette and the distance between an erythrocyte trapped and in the microvessel lumen and the occluding probe is measured as a function of time. The microvessel diameter is also measured. Volume flux (J,JS) is estimated using the initial erythrocyte velocity. Hydraulic conductivity (Lp) is determined as the slope of J,JS as a function of mean occluded intraluminal pressure (P).



Figure 3.

The isolated venule technique. (A) Schematic of the experimental setup. A venule is dissected, excised, and mounted on two glass micropipettes and bathed with an albumin physiological saline solution at 37°C. Pressure and flow are controlled by altering the heights of the inflow and outflow reservoirs, which are mounted on manometers. (B) To assess permeability, the fluorescence intensity of a window containing the venule and the nearby extraluminal area is measured. The inflow is quickly switched from normal APSS to APSS containing a fluorochrome‐labeled tracer (e.g. FITC‐albumin), causing a step increase fluorescence intensity (ΔIfo) that is proportional to the number of solute tracer molecules that have entered the lumen. This is followed by a gradual increase in intensity (dIf/dt)o. The rate of (dIf/dt)o is proportional to solute transport across the microvascular wall. The intensity returns to the basal level when the tracer is washed out from the lumen.

Adapted from 34 and 79 with permission


Figure 4.

Cultured endothelial cell (EC) permeability models. An EC monolayer is grown on a porous membrane that is placed between two chambers. (A) A vertical chamber system is shown, in which the upper chamber represents the “luminal” side and the tower chamber represents the “abluminal” side of the endothelial monolayer. Note that the fluid levels of the upper and lower chambers are kept at the same height to minimize any hydrostatic differences between the two chambers. (B) A horizontal setup.



Figure 5.

Ultrastructure of canine cardiac postcapillary venule. The electron micrograph demonstrates the complex morphology of endothelial cells (ECs). Possible transport pathways for solutes are indicated by different arrows. Caveolae and vesicles are abundant throughout the EC. WP. Weibel—Palade body. Durán, WN and Berendsen P (unpublished). (See page 3 in colour section at the back of the book)



Figure 6.

Routes of fluid and solute transport across the endothelium. Two pathways, one paracellular and one transcellular, have been proposed. In the paracellular pathway, fluid and solutes pass between endothelial cells (ECs), with the major barrier being the tightness of junctions between ECs. Several molecules thought to regulate junctional integrity are shown. In the transcellular pathway, active vesicular transport, or the formation of channels of VVOs, is proposed as a route for fluid and solute passage.



Figure 7.

Signal transduction pathways in microvascular permeability. Receptors are activated upon binding to their respective agonists and interact with downstream signaling pathways. The diagram reflects a typical set of cascades activated by permeability agonists; VEGF is used as an example.



Figure 8.

Transfection of ca‐ROCK protein increases coronary venular permeability. Isolated porcine coronary venules were transfected with 3 μg/ml ca‐ROCK protein for 120min in absence (solid line, squares, n = 6) or presence (dashed line, triangles, n = 4) of the ROCK inhibitor Y‐27632 (5 × 10−6M). Y‐27632 was added to the bath 20min prior to the addition of ca‐ROCK. Transfection was initiated at 0min and continued throughout the time course. *Indicates P < 0.05 vs. baseline permeability (0‐min time point). ‡Indicates P < 0.05 vs. ca‐ROCK transfection when Y‐27632 is present. From Ref. 307 with permission.



Figure 9.

MLC phosphorylation as a mechanism of endothelial barrier regulation. Inflammatory stimuli cause elevated phosphorylation of MLC on Thr‐18/Ser‐19 in endothelial cells, which in turn elevates actin—myosin mediated contraction. This leads to increased cellular centrifugal force that promotes opening of endothelial cell—cell junctions, causing increased endothelial permeability. MLC phosphorylation increases when MLCK activity is elevated or MLCP activity is reduced. The Rho/ROCK pathway can inactivate MLCP through phosphorylation of the MLCP targeting subunit MYPT‐1 on Thr‐696. (See page 4 in colour section at the back of the book)



Figure 10.

ROCK‐mediated tension and barrier dysfunction in coronary venular endothelial cell (EC) monolayers. Transfection of ca‐ROCK (3 μg/ml) into cultured coronary venular EC (CVEC) monolayers increases isometric tension (triangles) and causes a drop in TER (solid line). A time course overlay of ca‐ROCK‐stimulated changes in TER and tension shows that these two events occur simultaneously. From Ref. 307 with permission.



Figure 11.

Adherens junctions in endothelial cells (EC). (A) Adhesion between ECs is achieved by homotypic binding of VE‐cadherin molecules at endothelial cell—cell junctions. Several catenins (p120, α, β, γ) are associated with VE‐cadherin and regulate its homotypic binding, as well as linkage to the cytoskeleton. (B) Under stimulated conditions, such as during VEGF treatment, VE‐cadherin, β‐catenin, and p120 catenin are phosphorylated on tyrosine residues, and a reorganization of endothelial cell—cell junctions occurs, characterized by the appearance of several VE‐cadherin rich finger‐like projections (arrows). Scale bar represents 20μm. Images from Ref. 207 with permission. (See page 4 in colour section at the back of the book)



Figure 12.

PAF‐induced NO production. (A) NO concentration was measured in the suffusate of the hamster cheek pouch using a Nitric Oxide Analyzer 407. (B) Perivenular NO concentration was determined using a NO‐sensitive electrode in the mouse cremaster muscle. NO concentration rose quickly upon topical administration of PAF in both tissues. “From 407 with permission.



Figure 13.

Obligatory role of eNOS‐mediated NO production in PAF‐induced hyperpermeability in vivo. PAF‐induced hyperpermeability is markedly reduced in eNOS−/−mice compared to wild‐type control mice. Typical intravital fluorescence microscopy images of the respective cremaster muscles are shown below the IOI graph.

Adapted from Refs 74 and 410 with permission. (See page 4 in colour section at the back of the book)


Figure 14.

Translocation of eNOS. Top Panel: (A) Isolation of lipid rafts domains was done in control, Ach, or PAF‐treated cells. Fractions were probed against caveolin‐1 and eNOS. (A) ACh treatment. (B) PAF treatment. The blots represent three independent experiments. Bottom Panel: The influence of PAF and ACh on the distribution of eNOS‐GFP was assessed by fluorescence of GFP. The images are representative of three independent experiments. (A) ECV304‐eNOS‐GFP. (B) Human dermal microvascular cells stained with anti‐eNOS (red) and anti‐caveolin‐1 (green) antibodies. From Ref. 434 with permission. (See page 5 in colour section at the back of the book)



Figure 15.

Diagram of eNOS translocation process and eNOS activation. (A) Shows a simplified diagram of the eNOS translocation hypothesis. eNOS is anchored in caveolae through myristoylation and palmitoylation, and is associated with caveolin‐1. Upon stimulation with agonists, eNOS becomes phosphorylated, dissociates from caveolin‐1, and translocates from the caveolae to either the Golgi apparatus or the cytosol. It is postulated that preferential translocation to different compartments is related to vascular function and may allow eNOS to interact with specific function‐related target molecules. Even though the drawing depicts eNOS as being “free” in the cytosol, our recent‐unpublished data indicate that eNOS may be internalized to its intracellular targets via caveolae. (B). Several signals can regulate eNOS activity. While association with caveolin‐1 and phosphorylation of Thr‐497 decrease enzyme activity (NO production), interaction with Ca2+‐calmodulin and HSP90, phosphorylation on Ser‐1177 by Akt, and PKC activation can all increase activity. Panel A redrawn from 410 with permission.



Figure 16.

PAF‐induced eNOS phosphorylation and hyperpermeability in ECV304‐eNOSGFP cells. (A) ACh and PAF activate eNOS by Ser‐1177 phosphorylation and Thr495 dephosphorylation. (B) Inhibition of muscarinic ACh and PAF receptors blocks Ser‐1177 phosphorylation β‐actin served as a (loading) control in experiments shown in (A) and (B). (C) PAF increases permeability in ECVeNOS‐GFP cells. Monolayers of ECVeNOS‐GFP cells were treated with PAF and permeability to FITC‐Dx‐70 was measured. Data are expressed as permeability coefficients (mean ± SEM) for control and PAF‐stimulated monolayers. *P < 0.05, n = 6. From 434 with permission.



Figure 17.

Functions of Rap‐1. The diagram shows known and putative effectors and targets of Rap‐1. Arrows indicate stimulation, blunt ends denote inhibition.

Adapted from Ref. 457 with permission
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Article Title: Microcirculatory Exchange Function
 

How to Cite

Walter N. Durán, Fabiola A. Sánchez, Jerome W. Breslin. Microcirculatory Exchange Function. Compr Physiol 2011, Supplement 9: Handbook of Physiology, The Cardiovascular System, Microcirculation: 81-124. First published in print 2008. doi: 10.1002/cphy.cp020404