Vascular Endothelial Cell Behavior in Complex Mechanical

Subscriber access provided by University of Sunderland

Review

Vascular endothelial cell behavior in complex mechanical microenvironments Bryan James, and Josephine Allen ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.8b00628 • Publication Date (Web): 22 Oct 2018 Downloaded from http://pubs.acs.org on October 27, 2018

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Biomaterials Science & Engineering

Vascular endothelial cell behavior in complex mechanical microenvironments

Bryan D. James‡,∥ and Josephine Allen*,‡,§,

‡Department

of Materials Science & Engineering, University of Florida, 100 Rhines Hall, PO Box

116400, Gainesville, FL 32611, United States of America

§Institute for Cell and Tissue Science and Engineering, 300 Weil Hall, PO Box 116550, Gainesville,

FL 32611, United States of America

∥Institute

for Computational Engineering, University of Florida, 300 Weil Hall, PO Box 116550,

Gainesville, FL 32611, United States of America

*[email protected]

ACS Paragon Plus Environment

ACS Biomaterials Science & Engineering 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Abstract

The vascular mechanical microenvironment consists of a mixture of spatially and temporally changing mechanical forces. This exposes vascular endothelial cells to both hemodynamic forces (fluid flow, cyclic stretching, lateral pressure) and vessel forces (basement membrane mechanical and topographical properties). The vascular mechanical microenvironment is “complex” because these forces are dynamic and interrelated. Endothelial cells sense these forces through mechanosensory structures and transduce them into functional responses via mechanotransduction pathways culminating in behavior directly affecting vascular health. Recent in vitro studies have shown that endothelial cells respond in nuanced and unique ways to combinations of hemodynamic and vessel forces as compared to any single mechanical force. Understanding the interactive effects of the complex mechanical microenvironment on vascular endothelial behavior offers the opportunity to design future biomaterials and biomedical devices from the bottom-up by engineering for the cellular response. This review intends to describe and define 1) the blood vessel structure; 2) the complex mechanical microenvironment of the vascular endothelium; 3) the process in which vascular endothelial cells sense mechanical forces; and 4) the effect of mechanical forces on vascular endothelial cells with specific attention to recent works investigating the influence of combinations of mechanical forces. We conclude this review by providing our perspective on how the field can move forward to elucidate the effects of the complex mechanical microenvironment on vascular endothelial cell behavior.

Keywords mechanotransduction, shear stress, cyclic stretch, stiffness, topography, pressure


Page 2 of 85

Page 3 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Introduction Within the vasculature, a collection of mechanical forces acts on the blood vessel. These forces are hemodynamic-derived (related to blood flow) and vessel-derived (related to the vessel wall). The hemodynamic forces include: fluid shear stresses from flowing blood, cyclic stretching from the expansion and contraction of the blood vessel, and the lateral pressure from the blood. The vessel forces include: the mechanical and topographical properties of the vessel wall. These forces together compose the complex mechanical microenvironment of the endothelium. In this review, we consider and refer to the vascular mechanical microenvironment as “complex” because the mechanical forces present are dynamic and interrelated. More and more, the mechanical component of the cellular microenvironment is being recognized for participating in the regulation of cell phenotype.1–3 Cells respond to their mechanical microenvironment through the process of mechanotransduction, in which the cell senses a mechanical force and translates it into a biochemical signal.4 In a seminal study, the stiffness of the underlying substrate was shown to mediate mesenchymal stem cell differentiation5. Building on this, other studies have shown the effect of not only the elastic (time-independent), but also the viscoelastic (time-dependent) mechanical properties of the substrate material.6,7 Moreover, mesenchymal stem cells were shown to have a mechanical memory, in which they displayed an apparent difference in differentiation depending on the history of their mechanical environment.8 In fact, phenomenological correlations have been formulated relating individual mechanical forces, such as fluid shear stress, to endothelial cell phenotype and cardiovascular disease susceptibility. For example, endothelial cells express an atherogenic phenotype (being susceptible to atherosclerosis) and an atheroprotective phenotype (being inhibitory to atherosclerosis) depending on the fluid dynamics of the blood vessel.9 These two phenotypes are generally characterized by the up or down regulation of



inflammatory genes and the permeability of the endothelium to low-density lipoproteins.10 Increasingly, the complexity of the mechanical microenvironment is being appreciated for its contribution to cellular behavior.

Cardiovascular disease occurs and progresses with changes in the complex mechanical microenvironment. Hypertension results in not only changes in the pressure pulse amplitude acting on the blood vessel wall, but also increases the magnitude of the vessel stretching because the outward force (pressure) acting on the wall is greater.11 Likewise, atherosclerosis leads to calcification of the blood vessel and the formation of an atherosclerotic plaques, which contribute to changes in the local mechanical properties and fluid shear stresses around the plaque.12–15 These changes are not of individual forces, but are rather of combinations of forces. They are not static and global but are rather dynamic and local. In this way, diseased states exist having their own characteristic complex mechanical microenvironment.

This review intends to describe the structure of the vasculature, to define the vascular endothelial cell complex mechanical microenvironment that exists in both arteries and veins, to describe the process in which vascular endothelial cells sense mechanical forces, and to describe the body of knowledge learned from in vitro studies on the influence of mechanical forces on vascular endothelial cell phenotype. We give significant focus to several recent studies looking at the combined effect of mechanical forces on vascular endothelial cells, which have elicited responses unique to these more complex mechanical microenvironments. Lastly, we provide our perspective on how the field can move forward with continued research efforts aimed at elucidating the dynamic cellular response resulting from mechanical microenvironments.


Page 4 of 85

Page 5 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Vasculature Structure and Physiology

To understand the complex mechanical microenvironment requires a discussion on the structure of a typical blood vessel because the mechanical forces are largely dependent on the structure of the vessel. Changes in the vessel’s composition affects the mechanical properties and the mechanical response of the vessel which affect the behavior of the endothelium. The vascular system relies on an extensively branched closed-network of arteries, veins, and capillaries to pump blood to every region of the body. Although each vessel type has a slightly different structure, but in general arteries and veins consist of three concentric layers: 1) the tunica intima, 2) the tunica media, and 3) the tunica adventitia (Figure 1A).

Figure 1. A histological depiction of the arterial intima, media, and adventitia in a human coronary artery; dashed lines represent the internal and external elastic lamina (A). The blood vessel has a complex architecture consisting of concentric layers composed of an assortment of extracellular



matrix proteins, glycoproteins, proteoglycans, and growth factors specific to each layer (B). Diffuse intimal thickening occurs within the first few decades of life preceding atherosclerotic plaque formation. Adapted with permission from ref 16. Copyright 2016 Portland Press Ltd.

Intimal Layer

The tunica intima is the innermost layer of the blood vessel; composed of a monolayer of endothelial cells (endothelium) supported by a thin, proteinaceous basement membrane (Figure 1B).17 The endothelium serves as a blood barrier and regulates nutrient exchange from the blood to the surrounding tissue. The endothelial lining varies with blood vessel location in the body. In most tissue, the lining is continuous; however, it can be fenestrated, such as in endocrine and exocrine glands, or even discontinuous in the liver, the spleen, and the bone marrow allowing direct blood contact with subendothelial layers.17 These different arrangements of the endothelium are due to differing levels of hormone and metabolite exchange required at site specific instances of the vascularized tissue.17

One of the important cellular structures related to this is the glycocalyx, a continuously secreted membrane bound layer of proteoglycans, glycoproteins, and solute molecules. This layer extends from the endothelial cell surface; however, measurement of the glycocalyx’s thickness is method dependent with reported thicknesses ranging from nanometers to micrometers.18,19 Proteoglycans within the glycocalyx layer are graft co-polymers consisting of a protein core with glycosaminoglycan chains. The core proteins can be syndecan, glypican, perlecan, versican,


Page 6 of 85

Page 7 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


decorin, biglycan, and mimecan. The glycosaminoglycan content is primarily heparan sulfate, followed by chondroitin sulfate, dermatan sulfate, and keratan sulfate. Hyaluronic acid is a long polymeric molecule and is the only glycosaminoglycan that is unbound to a core protein. Specific combinations of core proteins and glycosaminoglycans exist with various functional significance.20,21 The proteoglycan composition is integral to the function of the glycocalyx layer and removal of any component drastically affects its properties.20 The glycoproteins of the glycocalyx are terminated with sialic acids, which play an important role in processes such as adhesion, coagulation, and fibrinolysis.20 The glycocalyx has been called the gatekeeper of the endothelium because it functions as a barrier to vascular permeability, as a mediator of erythrocyte, platelet, and neutrophil interactions, and as a reservoir of soluble factors.19,20,22 The glycocalyx layer is continuously removed by fluid flow and its products regulate the underlying smooth muscle cell phenotype.18,23 Additionally, vascular permeability is affected by the integrity of tight junctions and adherens junctions as well as transcellular pathways (vesicular trafficking).24,25 The endothelium/glycocalyx barrier is susceptible to changes in hemodynamics that accompany diseased states. Specifically, diseased states disrupt the integrity of the endothelium lining leading to leaky junctions, gaps between the cells, and increasing the permeability of the endothelium.24,26

Increased permeability of the endothelium may facilitate the migration of low-density lipoproteins (LDL) from the blood into the intimal layer.27 Accumulation of LDL in the intima and the LDL’s subsequent oxidation initiates the onset of atherosclerosis. The oxidation of LDL incites an inflammatory response by the endothelium, which recruits circulating monocytes from the blood. P-selectin and vascular cell adhesion molecule 1 are expressed and chemokines are produced by the activated endothelial cells facilitating monocyte tethering and rolling.28,29 Upon recruitment,



monocytes migrate below the endothelial monolayer into the intima, which is promoted by the activated endothelial cells displaying intercellular adhesion molecule 1 and producing chemokine gradients.10,30,31 In the intima, the monocytes then differentiate into macrophages and consume the oxidized LDL becoming foam cells (lipid-loaded cells). Neighboring smooth muscle cells in the medial layer are then targeted by secreted chemokines and growth factors from the differentiated macrophages and activated endothelium. These smooth muscle cells proliferate and synthesize extracellular matrix forming the body of an atherosclerotic plaque. Macrophage foam cells undergo apoptosis leading to the formation of the lipid-rich necrotic core of the plaque. The arterial vessel diameter narrows with the growing plaque. Secondary cell death from insufficient clearing of cellular debris prompts smooth muscle cell death in the plaque. At the edge of the plaque, a collection of inflammatory cells enzymatically breakdown the plaque extracellular matrix material. This degradation makes the plaque vulnerable to rupture.31,32 The plaque may either continue to grow narrowing the vessel diameter, incite endothelial cell death leading to thrombus formation, or rupture releasing highly thrombotic material into the bloodstream potentiating ischemia and infarction of downstream tissue.32 Endothelial cell dysfunction is a key instigator of atherosclerosis occurrence and progression.

Underlying the endothelium is the basement membrane, which serves as both a support structure for endothelial cell anchorage and a border between the endothelium and the surrounding vascular connective tissue.17 Important to the functionality of the basement membrane is its composition, which contains type IV, XV, and XVIII collagens, laminin, nidogen, perlecan, heparan sulfate, and von Willebrand-factor (Figure 1B).17,33 Type IV collagen within the basement membrane forms a two-dimensional network with a sheet structure and chicken wire-like appearance


Page 8 of 85

Page 9 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


contributing to the mechanical stability of the membrane.17 Laminins form an independent network and are necessary for basement membrane formation.17,34–37 The laminin network contains several cell adhesion receptors, such as integrins and sydecans.17,38,39 Nidogens, a type of sulfated glycoprotein, act to connect the type IV collagen network and laminin network.17,40,41 Type XVIII collagen is suspected to be involved in anchoring the basement membrane to the surrounding connective tissue.17,42 Perlecan is the major heparan sulfate proteoglycan in the basement membrane and serves multiple functions. It acts to stabilize the basement membrane structure by preventing degradation of laminin.17,43–46 Additionally, perlecan binds low density lipoprotein and interacts with fibroblast growth factor, vascular endothelial growth factor, and platelet-derived growth factor.17,47 Von Willebrand factor is a blood glycoprotein that plays a key role in hemostasis. It is embedded in the membrane, a specific feature of the endothelial cell basement membrane.17,48,49 The diseased state of the intima is associated with the deposition of fibronectin and type I and III collagens.50–53 The tunica intima is home to the endothelium, which serves as the gatekeeper between the blood and the surrounding tissue.

Medial Layer

The tunica media is the middle layer of the blood vessel; composed primarily of a thick collagenand elastin- rich matrix with embedded mural cells (Figure 1B).17 In arteries and veins vascular smooth muscle cells serve to regulate vascular tone. Both extrinsic (global) and intrinsic (local) factors act to control smooth muscle cell regulation of vascular tone.54 One of the key extrinsic factors is endothelial cell-derived nitric oxide.54–61 These factors affect the dilation and constriction of the vessel, which is used to regulate blood flow. In healthy tissue, smooth muscle cells exist in



a quiescent, contractile phenotype whereas in diseased states, such as intimal hyperplasia or atherosclerosis, smooth muscle cells exhibit a proliferative, synthetic phenotype.15 Smooth muscle cell phenotype is largely dependent on the extracellular matrix, on growth factors and cytokines, and on physiology.17,62,63 Additionally, the phenotype is modulated by endothelial secreted products. In vitro co-culture studies of smooth muscle cells and endothelial cells have shown that a proliferating endothelium will stimulate proliferation of synthetic smooth muscle cells whereas a quiescent endothelium will inhibit their proliferation.23,64

The extracellular matrix enclosed within the medial layer contains elastin, fibrillin, fibulin, fibronectin, and type I and III collagens (Figure 1B).17,33 Elastin endows the blood vessel with its elasticity and resilience.17,65 Elastin is organized in lamella forming concentric layers with smooth muscle cells sandwiched in between them. Elastin and its degradation peptides also mitigate smooth muscle cell proliferation.17,33,66,67 In the event of vessel damage, elastic fibers are typically not replaced with new elastin, but instead are replaced by collagen leading to vessel stiffening.33 Additionally, the vessel may stiffen by calcification of the elastic lamellae.33 These changes in vessel mechanical properties will directly impact the characteristics of vessel expansion and contraction from pulsatile blood flow. Microfibrils, containing fibrillin-1 and fibulin-5 glycoproteins, connect concentric elastin lamellae aiding in their formation and integrity as well as in mediating cellular interactions.17,33,66,68 Fibronectin modifies the elasticity of the vessel and regulates the expression of other matrix proteins.33,69,70 Lastly, type I and III collagens give the vessel its tensile properties and set the limit on its expansibility.17 Whereas, elastin mitigated smooth muscle cell proliferation, soluble collagens stimulate it.15,71 In fact, the smooth muscle cell response is dependent on the type of collagen, its concentration, conformation and context within


Page 10 of 85

Page 11 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


the extracellular matrix.15 The tunica media largely provides the blood vessel its mechanical properties and its composition regulates smooth muscle cell phenotype.

Adventitial Layer

The tunica adventitia is the outermost layer of the blood vessel; composed of fibroelastic connective tissue (Figure 1B). Once thought to only contain nerves and fibroblasts, in recent years the adventitia has been shown to encompass a highly interactive community of embedded fibroblasts, myofibroblasts, macrophages, T cells, B cells, mast cells, dendritic cells, microvascular endothelial cells, pericytes, adipocytes, lymphocytes, and vascular progenitor cells.72–77 Two separate populations of progenitor cells exist with potential differentiation lineages being mural cells for one and macrophage-like cells for the other.72,78 Additionally, the adventitia participates in regulating vascular tone and vascular remodeling both of which will alter the endothelial cell mechanical environment.73,74 The collection of immune cells (macrophages, T cells, B cells, and mast cells) execute innate immune functions in response to foreign antigens.73,74,76,79 It is largely believed that the adventitia acts as the sensory structure for vessel damage and injury.76 The adventitia layer is largely collagenous, which gives the vessel its rupture properties. Specifically, it is composed of Type I and III collagens organized as fibrils and elastin. 14,15,80

Fibroblasts and myofibroblasts arrange longitudinally in the matrix. Together, they maintain

the fibrillar collagen of the layer.14 The tunica adventitia supports a community of cells and connects the vessel to the surrounding tissue.



Defining the Complex Mechanical Microenvironment

The vasculature produces a collection of dynamic mechanical forces, which form the complex mechanical microenvironment. These include hemodynamic forces (fluid flow, cyclic stretch, and lateral pressure) and vessel wall forces (the mechanical and topographical properties of the vessel wall) (Figure 2A).

Figure 2. The endothelium experiences a combination of interrelated hemodynamic forces from fluid flow, lateral pressure, cyclic stretch, and vessel wall forces from the mechanical (elastic and


Page 12 of 85

Page 13 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


viscoelastic) and topographical properties of the extracellular matrix. The hemodynamic forces are time-dependent, changing over the course of the cardiac cycle from the pumping action of the heart (B). The black dashed line represents the separation between systole and diastole. The stretching of the vessel lags the pressure pulse by a phase angle (difference between the purple and blue dashed lines), which is dependent on the viscoelastic properties of the vessel (D). The shear stress lags the cyclic stretch by the stress phase angle (difference between the blue and red dashed lines). The time-averaged wall shear stress (TAWSS), the oscillatory shear index (OSI), and the transverse wall shear stress (transWSS) are used to describe the different flow regimes in a blood vessel (C). The TAWSS is unable to describe changes in flow direction. The OSI can describe unidirectional reciprocating flows but is unable to detect multidirectional flows. The transWSS can discriminate and describe differences in multidirectional flows. The values presented for each are non-specific and are provided to illustrate the differences between the three flow regime descriptors. The vessel wall is a strain stiffening viscoelastic body; therefore, its stress-strain behavior is both time-dependent (creep and stress-relaxation) and strain-dependent (D).

Fluid Flow

Blood flow varies spatially and temporally throughout the vasculature. Flows in the vasculature are oscillatory, pulsatile and differ between straight and bifurcating blood vessels, the result of the pulsatile pumping action of the heart over the cardiac cycle.81 In straight blood vessels, flow is laminar and largely unidirectional while at regions of blood vessel curvature it is disturbed and bidirectional. Straight blood vessels also exhibit greater wall shear stresses.81 Disturbed flows



arising at bifurcating and curving blood vessels display flow separation with transient flow reversals, lower average wall shear stresses, and occasional turbulence.81 As a result of these changing flow patterns, the likelihood of atherosclerotic plaques forming at bifurcations is high compared to straight vessels.81–83

Wall shear stress is a transient vector quantity, which throughout the body has a magnitude ranging from 0–20 Pa (0–200 dynes/cm2).84–87 In the straight sections of arteries, the average wall shear stress magnitude is maintained at ~1.5 Pa (15 dynes/cm2). Generally speaking, the average wall shear stress magnitude increases with increasing distance from the heart for both arteries and veins.88 The measured average wall shear stress: in the aorta is 0.448 Pa (4.48 dynes/cm2), in the carotid artery is 0.640 Pa (6.40 dynes/cm2), in the femoral artery is 0.734 Pa (7.34 dynes/cm2), in small arteries is 3.200 Pa (32.00 dynes/cm2), in large veins is 0.524 Pa (5.24 dynes/cm2), in small veins is 1.080 Pa (10.80 dynes/cm2), and in the vena cava is 0.300 Pa (3.00 dynes/cm2).88 Moreover, flow changes over the course of the cardiac cycle and consequently, so does the wall shear stress, making it a transient quantity (Figure 2B). Because of this, more than a simple arithmetic average is necessary to describe the wall shear stress. Hemodynamic properties in the vasculature can be quantified in several different ways over a period of interest, typically expressed as the cardiac cycle. The magnitude of the wall shear stress can be described by either the maximum value or the time-averaged value (TAWSS) over the period of interest. The directionality of the wall shear stress can be expressed as the direction of the wall shear stress vector, but more descriptive quantities are the oscillatory shear index (OSI) and the transverse wall shear stress (transWSS).89,90


Page 14 of 85

Page 15 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


1 𝑇

𝑇𝐴𝑊𝑆𝑆 = 𝑇∫0|𝝉𝑾𝑺𝑺|𝑑𝑡

𝑂𝑆𝐼 = 1 ―

|∫𝑇0𝝉𝐖𝐒𝐒𝑑𝑡| 𝑇 ∫0|𝝉𝐖𝐒𝐒|𝑑𝑡

𝑡𝑟𝑎𝑛𝑠𝑊𝑆𝑆 =

1 𝑇 𝑇∫ 0

|

(1)

= 1 ―

(

|𝝉𝐖𝐒𝐒|

(2)

𝑇𝐴𝑊𝑆𝑆

𝝉𝐖𝐒𝐒 ∙ 𝒏 ×

𝑇

∫0𝝉𝐖𝐒𝐒𝑑𝑡

|∫𝑇0𝝉𝐖𝐒𝐒𝑑𝑡|

)|

𝑑𝑡

(3)

In equations 1-3, 𝝉𝐖𝐒𝐒 is the wall shear stress vector, T is the time period of interest, and 𝒏 is the unit normal vector to the wall. The OSI was formulated to characterize the extent that wall shear stress reverses direction. An OSI of 0 indicates perfectly unidirectional flow while an OSI of 1 indicates complete flow reversal.89,90 Whereas, the OSI characterizes flow reversal, the transWSS characterizes the extent to which the wall shear stress acts in the direction perpendicular to the primary flow direction. Other metrics have been proposed, but have all been found to capture the same features as those of the TAWSS, OSI, and transWSS or combination of the three (Figure 2C).86,90–99 In general, wall shear stresses are transient vector quantities with location-specific magnitudes ranging between 0–20 Pa (0–200 dynes/cm2).

Cyclic Stretch

The vessel wall expands and contracts in response to lateral pressure generating a cyclic stretch, also known as circumferential strain, varying spatially and temporally throughout the vasculature. The mechanical response of the blood vessel due to the lateral pressure acting on the vessel should not be confused with vasomotion and vascular tone, both referring to the extent to which smooth



muscle cells change the diameter of the blood vessel from its relaxed state (basal tone) to regulate blood flow.54 Similar to fluid flow, circumferential strain of the vessel wall is either uni- or biaxial and changes between straight and bifurcating vessels.100 The stretching magnitude oscillates between the systolic and diastolic blood pressure with the frequency of the cardiac cycle (Figure 2B). The strains are maximum at systole and vary with vessel type. At systole, the strain is 9–12% for the aorta, 6–10% for the pulmonary arteries, 1–2% for the carotid arteries, and 2–15% for the femoral arteries.11,101–110 Conversely, lateral strains are about 1% being restricted due to the surrounding tissue.101 As with blood flow patterns, diseased states such as hypertension and hypotension also impact cyclic stretching by deviations from the healthy strain as well as the frequency of stretch due to elevated heart rate.100 Under such conditions, strains can be as high as 20% as seen in hypertension.111 The vessel wall response to the pressure in the vessel with each cardiac cycle lags by a phase angle. In large diameter vessels, the phase angle is between 5–10 degrees (Figure 2B). The lag is due to the viscoelastic mechanical properties of the vessel.101 The dependence of cyclic stretching on hemodynamics intimately couples the two forces. The stretching of the vessel can be simulated using computational fluid dynamics simulations coupled with finite element simulations of vessel mechanics termed fluid-structure interaction simulations. These have shown that moving vessel walls in healthy femoral and carotid arteries reduce the wall shear stress by 5–25%.87,112–114 Conversely, simulations of abdominal aortic aneurysm have shown that flow has little effect on the tissue stresses.87,115,116 One parameter relating the two factors is the temporal phase angle between the wall shear stress and the cyclic stretch, termed the stress phase angle (Figure 2B). It has been shown that atherosclerotic plaques localize to places where the two are most out-of-phase.117 Much like with fluid flow, blood vessel cyclic stretch is a transient quantity with location-specificity and a magnitude ranging between 1–20%.


Page 16 of 85

Page 17 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Lateral Pressure

The lateral pressure is a linear combination of several pressure terms varying spatially and temporally throughout the vasculature. The total pressure is composed of an ambient pressure term, a hydrostatic pressure term, a dynamic pressure term, and a potential energy pressure term.101 The ambient, hydrostatic, and potential energy pressure terms represent the lateral pressure acting on the vessel. The ambient pressure term is typically taken as 0 Pa. The hydrostatic pressure term is taken as 0 Pa at the tricuspid valve of the heart and changes as a function of vertical displacement with reference to that position.101 In an upright position, the hydrostatic pressure experienced in the head is subatmospheric (-1.3 kPa or -10 mmHg) and in the feet is superatmospheric (12 kPa or 90 mmHg).118 These changes of pressure are regulated by the venous pump system of the vasculature and only exhibit these extremes when the body is stationary.118 The dynamic term drives blood flow and does not play a role in the stretching of the vessel. The final term, the potential energy pressure, is generated by the pumping action of the heart and is consequently pulsatile (Figure 2B). It oscillates between the systolic and diastolic blood pressures centered at a time-averaged value.101 This implies that the lateral pressure is dependent on body posture, activity level, age, vascular tone, and disease.101,104,119 As a result, blood pressure as measured by a sphygmomanometer is proportional to the lateral pressure, but not equal to it.101 Typical measured blood pressures in the aorta are 16 kPa (120 mmHg) for the systole and 10.6 kPa (80 mmHg) for the diastole. The time-average pressure decreases with greater distance from the heart, dropping to 12.6 kPa (95 mmHg ) in large arteries and drastically to 4 kPa (30 mmHg) at the entrance to capillaries.101,120 In venous networks, the time-average pressure is much lower being 1.3–1.6 kPa (10–12 mmHg) and decreases slowly to zero at the right atrium of the heart.101,120 Hypertension is



defined by four ranges of increased systolic and diastolic blood pressures as measured at the aorta: elevated, stage 1, stage 2, and crisis. The time-averaged blood pressure ranges for each are: 13.3– 13.9 kPa (100–105 mmHg), 14.0–15.1 kPa (110–114 mmHg), 15.3 kPa (>115 mmHg), and 20.0 kPa (>150 mmHg).121 As with the other hemodynamic forces, the lateral pressure is a transient quantity with location-specificity and a magnitude ranging from 1.3–16 kPa (10–120 mmHg).

Basement Membrane

The intimal layer basement membrane is a compliant network of densely packed proteinaceous fibers for supporting endothelial cells (Figure 1B).17,80,122 The elastic modulus of the membrane ranges from 8–100 kPa.17,80,122 Specifically, measurement of the saphenous vein elastic modulus by microindentation was 8.2 kPa.123 Measurement by microindentation of the elastic modulus for the outermost region of the intima of a porcine artery was 69.0 kPa.124 Conversely, measurements by tensile testing have found the elastic modulus to be approximately 1–2 orders of magnitude greater than those measured by indentation methods.125–127 This measurement discrepancy is attributed to the nature of each technique in that tensile testing is a macroscopic, global measurement while indentation is a microscopic, local measurement of the mechanical properties.127,128 The length scale of the component is important as well when considering its stiffness. In general, the elastic modulus decreases with increasing length scale.128 In comparison to the elastic modulus of tissue being on the order of kilopascals, the elastic modulus of fibrillar collagen is on the order of megapascals, and the elastic modulus of a collagen molecule is on the order of gigapascals.129–133 Cells exist on the microstructural length scale and thus experience the tissue mechanical properties.6,128 The extracellular matrix also changes with diseased states.


Page 18 of 85

Page 19 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Atherosclerosis is associated with arterial calcification, which elevates the extracellular matrix stiffness by approximately 5 to 8 times.12–15 It is important to not only consider the elastic response but also the viscoelastic response (Figure 2D). Using atomic force microscopy, the viscoelastic response of human femoral arteries was measured and modeled as a three element standard linear solid yielding a strain relaxation time of 16.9 seconds and a stress relaxation time of 29.3 seconds.134 The viscoelastic properties of elastin meshes derived from porcine thoracic aortas showed a strong dependence on hydration becoming more viscous and less anisotropic with water loss.135 Collagen gel storage and loss modulus both decreased with the addition of increasing concentrations of fibronectin and laminin into the network. This change in collagen gel properties was attributed to the formation of a looser network structure as a result of the added proteins.136 This highlights the importance of each component to the overall mechanical properties of the membrane. Additionally, it is well known that biopolymers exhibit strain stiffening behavior, a phenomenon that has been shown to be true for the basement membrane, as well (Figure 2D).129,137,138 This strain-stiffening behavior has significant implications as the basement membrane is deformed in a non-linear transient manner over the cardiac cycle. Morphologically, the basement membrane has the appearance of a felt-like mesh work with nano- to microtopographical features derived from pores, fibers, and elevations.122,139,140 As studied in the rhesus macaque, the vascular basement membrane consists of pore sizes between 59–63 nm in diameter and fibers between 30–31 nm in diameter.141 Also, the thickness of the basement membrane varies with vessel type: the aorta being the largest (~500 nm) and decreasing for the carotid artery (~350 nm) and more so for the saphenous vein (~100 nm).141 Overall, it can be said that the intimal layer basement membrane is a mesh with submicron topographical features and expresses dynamic mechanical properties over the course of the cardiac cycle.



Figure 3. The hemodynamic forces vary spatially throughout the body and change with the onset and progression of disease states. Typically, in the diseased state (hypertension/atherosclerosis), the time-averaged fluid wall shear stress (WSS) decreases (↓), the magnitude of cyclic stretching (CS) decreases (↓), and the time-averaged lateral pressure (LP) increases (↑). The values presented


Page 20 of 85

Page 21 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


are not absolutes and vary person to person and with the arterial geometry. References: a – 88 ,b – 120,

c – 142, d – 102, e – 143, f – 144, g – 145, h – 146, i – 147, j – 148, k – 149, l – 150.

The vascular complex mechanical microenvironment is both spatially and temporally diverse. Each hemodynamic force originates from the pumping action of the heart and each interacts with one another (Figure 2B). These forces are location-specific and change with diseased state (Figure 3). Pulsatile lateral pressure stresses the vessel wall causing it to expand and contract, which is accompanied by a fluid wall shear stress propelled by the pressure gradient in the vessel. The cyclic stretching likewise dynamically strains the basement membrane in a non-linear manner deforming the microstructure and deviating the basement membrane from its static mechanical properties. The vessel mechanical properties in turn contribute to the extent of circumferential strain and vascular tone regulates the fluid flow. The mechanical forces of the vasculature are local and complex, which collectively act on the endothelium. It is important that these factors are welldefined before we move the discussion to the way the endothelium is affected by these forces.

Sensing the Mechanical Microenvironment

The emerging field of mechanobiology studies the processes by which cells detect and respond to their mechanical microenvironment. Physical forces are sensed by cellular structures interacting with the environment (mechanosensing) and are transduced into biochemical signals via signal transduction pathways (mechanotransduction). This general action is the mechanism by which the endothelium engages with mechanical forces. The study of mechanotransduction by the endothelium is on its own the subject of numerous texts.101,151–153 The following paragraphs are to



provide the reader with an overview of the different mechanosensors operating in the endothelial cell to offer a general understanding of the many structures that allow endothelial cells to respond to their complex mechanical microenvironment (Figure 4).

Figure 4. Endothelial cells sense their mechanical environment using numerous mechanosensory structures including: the plasma membrane, the nucleus, the glycocalyx, membrane-embedded Gprotein coupled receptors (GPCR) and G-proteins, membrane-embedded mechanically-activated ion channels, focal adhesions, and cell-cell junctions. These mechanosensors collectively form the endothelial cell mechanosensory system.

Plasma membrane

Mechanical deformation of the plasma membrane is theorized as a mechanism for activation of mechanosensitive structures. The plasma membrane is a dynamic phospholipid bilayer of variable


Page 22 of 85

Page 23 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


saturation, cholesterol content, and membrane bound proteins.154 Interactions between all three contribute to the membrane fluidity, the ability for lipids to move freely in the membrane.155 Because the membrane is exposed to applied forces either directly by acting on its surface or indirectly by acting on surface bound structures, it is theorized to be a mechanosensor.155 The mechanism for mechanosensitivity is hypothesized to be due to changes in membrane curvature, which changes both membrane fluidity and conformation of membrane bound proteins due to hydrophobic mismatch between the bound protein and the membrane. Collectively, these changes modify the lateral diffusion of membrane bound proteins. It has been shown that changes in membrane fluidity are able to discriminate between shear stress and stretch.156 Together these changes are believed to activate membrane bound ion channels, G-protein coupled receptors, or other changes.155

Nucleus

Cytoskeletal connections are capable of relaying mechanical forces directly to the nuclear membrane.157 Force transmission is achieved via LINC (linker of the nucleoskeleton and cytoskeleton) complexes connecting the cytoskeleton to the nucleoskeleton.158–160 It is through this coupling that mechanosensitive genes could be directly activated.160 In response to applied forces, the nucleus has been shown to deform and stiffen and is implicated in cell migration through threedimensional matrices.159,160 The proposed mechanism by which forces are sensed by the nucleus are numerous and are thought to be through stress-induced changes in protein conformation, translocation of transcriptional regulators, chromosome conformation and organization, and membrane dilation.161



Glycocalyx

It has been hypothesized that endothelial cells sense fluid shear with an intact glycocalyx through the extended surface of the glycosaminoglycan chains.18 Force is transmitted from the glycosaminoglycan chains through the membrane bound core proteins and subsequently distributed to mechanotransducing structures in the cell via the cytoskeleton. The core proteins have been shown to be sufficiently stiff to act as transmitters.162 In contrast, endothelial cells with a degraded glycocalyx sense shear directly at the plasma membrane surface.18 In such a situation, the force is still felt and balanced by restraining forces at adhesion sites. It has been suggested that the transmitted force at the adhesion sites is the same regardless of the integrity of the glycocalyx layer.18 Additionally, the heparan sulfate in the glycocalyx layer has been implicated in sensing hypotonic stress.163 The glycocalyx layer is increasingly being investigated for its role in mechanobiology and functional significance in medicine.164

G-proteins and G-protein coupled receptors

G-protein and G-protein coupled receptor activation processes are a potential membrane bound mechanosensor. In general, G-proteins and G-protein coupled receptors are plasma membrane proteins, which together act as a switch facilitating intracellular signaling.165,166 G-protein signaling proceeds when an extracellular ligand activates the transmembrane G-protein coupled receptor. This in turn causes the transfer of guanosine diphosphate (GDP) bound to the G-protein to exchange with G-protein coupled receptor bound guanosine triphosphate (GTP). Consequently, the heterotrimeric G-protein (composed of subunits α, β, γ) dissociates releasing the G subunit


Page 24 of 85

Page 25 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


with the newly bound GTP from the remaining peripheral membrane bound G subunits.165,166 Gproteins become activated within 1 second after mechanical stimulation.167–169 At times other than ligand activation, G-proteins and G-protein coupled receptors are free to move in the membrane without forming complexes.170,171 Because of this free movement, it is theorized that changes in the lateral diffusion of the G-protein induced by changes in either the plasma membrane fluidity, thickness, or polarity as a result of mechanical deformation would modify the total activation efficiency of the G-protein and G-protein coupled receptor complex.172,173 In this way, G-proteins and G-protein coupled receptor complexes are believed to be mechanosensory structures. Recently, dela Paz et al. investigated the influence of heparan sulfate proteoglycans on the mechanosensitive complex of platelet endothelial cell adhesion molecule-1 (PECAM-1) and heterotrimeric G protein subunit 𝐺𝛼𝑞/11, which is capable of discriminating between different flow profiles.174,175 It was suggested that the interaction between PECAM-1 and 𝐺𝛼𝑞/11 is mediated by the heparan sulfate of syndecan-1.174 G-proteins and G-protein coupled receptors are a potential mechanosensor due to their variable activation efficiency mediated by membrane deformation.

Mechanically-activated ion channels

Mechanically-activated ion channels are transmembrane proteins, which form pores to allow the selective passage of ions under applied stress. Two mechanisms are proposed for channel activation termed the ‘force-from-lipids’ models, which operates on the premise of membrane tension altering lipid-protein interactions and the ‘force-from-filaments’ model, which operates through interaction of the channel with direct connection to the extracellular matrix or cytoskeleton.176,177 Flow-activated ion channels allow the transfer of Na+, Ca2+, K+, and Cl- ions178.



It has been shown that shear stresses as low as 0.01 Pa (0.1 dyne/cm2) are capable of activating K+ channels whereas a shear stress of 0.03 Pa (0.3 dyne/cm2) is necessary to activate Cl- channels. Ion currents have been shown to saturate at 1-1.5 Pa (10-15 dyne/cm2) for K+ channels and at 0.35 Pa (3.5 dyne/cm2) for Cl- channels.179,180 Additionally, K+ and Cl- channel activation is sensitive to oscillatory flow frequency below a threshold frequency.179,180 Likewise, Cl- channel activation desensitizes under prolonged shear stress and rapidly above a threshold shear stress.178 Also, these channels activate independently and immediately upon the onset of flow.178 Because of the differential timing of the two channels (K+ and Cl-) and their variable responsiveness to flow conditions, it is theorized that endothelial cells use these two channels to discriminate between different types of flows.181 In recent years, the discovery of Piezo1 channels, mechanicallyactivated Na+ and Ca2+ ion channels, have been implicated in flow shear stress sensing.182–184 Rode et al. suggested that Piezo1 is required for elevated blood pressure during physical activity, but not sedentary blood pressure because Piezo1 responded to increased blood flow during exercise and activated vasoconstrictive action by smooth muscle cells.185,186 Stretch-activated channels allow the transfer of Na+, Ca2+, and K+ ions.178,187 Specifically, Ca2+ exchange by transient receptor potential vanilloid channel 4 is thought to participate in the mechanosensing of stretch due to its high expression and its activation in cyclically strained endothelial cells.11,188–190 Mechanicallyactivated ion channels are potential mechanosensors of fluid flow and cyclic stretch.

Adhesion sites

Cells detect and act on their extracellular matrix and cellular neighbors through adhesion sites. In general, cells probe their environment through three types of forces, traction forces, protrusive


Page 26 of 85

Page 27 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


forces, and cell-to-cell forces. Endothelial cells adhere to their extracellular matrix primarily with transmembrane integrin focal adhesions. Integrins are heterodimers consisting of single - and subunits joined into 24 known combinations.191 The different combinations bind to specific extracellular matrix proteins and adhesion molecules. The most characterized integrin binding ligand is the tripeptide alanine-glycine-aspartate (RGD), which is found in fibronectin, vitronectin, and other adhesion molecules.192 Integrins are a component of the greater adhesion structure known as a focal adhesion. These are multiprotein complexes, which support the connection between the adhesion ligand on the extracellular matrix protein to the actin cytoskeleton.193 Focal adhesions are dynamic structures that remodel in response to applied forces and strengthen upon pulling forces through their actin connections. It is believed that through the probing of the surface by these tractions forces, cells are able to detect the mechanical properties and topography of their extracellular matrix.4,194 In addition to sensing substrate properties, focal adhesion complexes are also involved in cellular migration. Cells migrate via the dynamic assembly and disassembly of focal adhesions and move in the direction of an increasing extracellular matrix gradient (i.e. an adhesion ligand density gradient), termed haptotaxis.195 An optimal ligand density exists for achieving maximum cell migration speed. In fact, under high ligand density cells are too firmly attached to migrate whereas under low ligand density cells have an abundance of binding sites.191,196 High lateral diffusion of integrins in the plasma membrane is associated with rapid migration whereas low diffusion is observed for stationary cells.197 Likewise, cells migrate up an increasing extracellular matrix rigidity gradient, termed durotaxis.198 However, it should be noted that the sensitivity to extracellular matrix rigidity plateaus above 300 kPa.191 Most recently, Gong et al. showed through analytical and Monte Carlo simulation that substrate viscosity stiffens soft substrates on a timescale faster than cellular traction forces, which enhances adhesion and



spreading whereas the viscous component of stiff substrates was shown to have no effect because of the plateau in sensitivity.199 Cells are also capable of distinguishing substrate thickness. Individual cells can sense an underlying stiff substrate through a soft gel below a critical thickness of ~10 µm. The critical thickness increases by an order of magnitude for cells in monolayer.200Additionally, cellular response to topography is termed, contact guidance.201 Numerous cell types have been shown to respond to topographic cues; for example, cells have been shown to align in the direction of grooves and also respond by differentiating into various lineages.202,203 Feature geometries can be derived from either native basement membrane architecture or engineered topographies.204 The mechanism of contact guidance is believed to be due to either variable protein adsorption to the topography, biasing of focal adhesion formation, or preferential actin polymerization from topography probing filopodia.203 Endothelial cells connect to and communicate with their neighbors through cell-cell junctions.205 The vascular endothelial (VE) cadherin-based adherens junction has been identified as a mechanosensor.206 These junctions are connected to the cytoskeleton and act to transmit mechanical forces between cells.207 This was first observed from fluid flow studies, which demonstrated that VE-cadherin was required in combination with vascular endothelial growth factor receptor 2 (VEGF-R2) and platelet endothelial cell adhesion molecule 1 (PECAM-1).208 Additionally, PECAM-1 specifically has been suggested as a mechanosensor by undergoing tyrosine phosphorylation upon application of forces in neighboring cells.11,209–211 Cellular adhesions sites to the extracellular matrix and to neighboring cells allow the cell to probe the material properties and surrounding environmental forces acting on it.


Page 28 of 85

Page 29 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


One question still to be answered is the process in which endothelial cells detect flow. Two theories attempt to describe the process, a decentralized theory and a bilayer membrane theory.155,212 The decentralized theory posits that mechanosensing structures detect the fluid shear stress and relay the force by means of the cytoskeleton to mechanotransducing structures across the cell.153,212 Conversely, the bilayer membrane theory, argues that the curvature and fluidity of the plasma membrane as altered by fluid shear stress is capable of activating mechanotransducing structures embedded in the membrane.155 Most recently, Dabagh et al. performed finite element simulations of a three-dimensional, multi-scale, multi-component, viscoelastic model of focally adhered endothelial cells exposed to oscillatory and multi-directional shear flows.213 The simulation results revealed that reversing and disturbed flows produce lower stresses due to the viscoelastic dampening of the endothelial cell structural components. Additionally, the forces felt at modeled mechanotranducers were found to be of enough magnitude to elicit activation. Taken together, it was proposed that the differential response by endothelial cells to atherogenic and atheroprotective flows could be due to a force threshold, which is overcome in unidirectional flows and missed in reversing flows. The difficulty in deducing the mechanism by which fluid flow is conferred into a biological response is due to the multiplex of sensing structures all being engaged seemingly at once. The challenge is discerning the order of engagement. Is there a primary sensing structure or is it a mechanosensory system composed of multiple mechanosensors? The commonality between both the decentralized and bilayer theories is that a plexus of mechanosensing structures are activated. Additionally, these structures are coincidentally engaged in sensing the other hemodynamic and vessel wall forces in combination with the fluid shear stress. The mechanical response of the cell to applied forces in general can be described by the tensegrity model, which is centralized around the cell existing in a state of tensile prestress through adhesion sites and the



cytoskeleton214. It offers a way to describe autonomous cellular shape stability under applied loads. In other words it can describe the shape change of the cell due to applied forces.214 The tensegrity model underlies both the decentralized theory and bilayer membrane theory of flow shear stress sensing. It can explain the ability of the cytoskeleton to relay and amplify mechanical signals to mechanotransducing structures across the cell and as a method for autonomous mechanical deformation of the cell membrane.

Figure 5. “Schematic diagram showing endothelial mechanotransduction and signaling induced by shear stress. Shear stress stimulates endothelial cells (ECs) through the activation of mechanosensors, including integrins, tyrosine kinase receptors (TKRs), G proteins and G-protein–


Page 30 of 85

Page 31 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


coupled receptors (GPCRs), ion channels, intercellular junction proteins (eg, vascular endothelial [VE]-cadherin and platelet/endothelial cell adhesion molecule 1 [PECAM-1]), caveolae, membrane lipids, and glycocalyx. These mechanosensors act through adaptor molecules (eg, SHC [Src homology 2 domain containing] transforming protein 1 [Shc]) to trigger the activation of signaling molecules such as Ras, Rho, phosphatidylinositol-3-kinase (PI3K), and mitogenactivated protein kinases (MAPKs), which then activate endothelial nitric oxide synthase (eNOS), Smad1/5, and the transcription factors and cofactors (eg, Kruppel-like factor 2 [KLF2], nuclear factor-κB [NF-κB], and activator protein 1 [AP-1]) to regulate the expression of several functional genes, such as eNOS, vascular cell adhesion protein 1 (VCAM-1), and monocyte chemoattractant protein-1 (MCP-1), as well as microRNAs (miRs). This diagram illustrates that multiple signaling pathways coordinate to form mechanoresponsive networks to modulate endothelial cell (EC) phenotype and function. AMPKs indicates AMP-activated protein kinases; BMPR, bone morphogenetic protein receptor; ERK, extracellular-signal-regulated kinase; FAK, focal adhesion kinase; HDAC, histone deacetylase; MEF2, myocyte enhancer factor-2; MPAKs, mitogenactivated protein kinases; mTOR, mammalian target of rapamycin; PYK2, proline-rich tyrosine kinase 2; SREBP2, sterol regulatory element binding protein 2; and VEGFR2, vascular endothelial growth factor receptor-2.” Figure and caption reproduced with permission from ref 215. Copyright 2014 Wolters Kluwer Health, Inc.

The downstream pathways activated by the mechanosensory system are numerous and complex, which continue to be discovered. The reader is directed to a collection of reviews and articles, which thoroughly describe the current understanding of endothelial mechanotransduction



signaling pathways for hemodynamic and vessel forces, the details of which are beyond the scope of this review.11,101,152,216 The most well studied in endothelial cells are the signaling pathways activated by fluid shear stress (Figure 5) and are shown to provide the reader with a subset of activated mechanotransduction pathways.9,153,212,215,217–224 Still much remains to be understood about the interactions of these pathways upon stimulation of combinations of mechanical forces.

There are numerous structures and proposed mechanisms by which endothelial cells sense their mechanical microenvironment. The primary sensing structure or collection of structures of a mechanosensory system remains to be determined. Currently, no single structure is known to sense the entire collection of forces making up the mechanical microenvironment. Each mechanosensor is connected in a network by the cytoskeleton and forces can be amplified from one sensor to another. Not only can forces be felt, but through cell-cell junctions, forces can be produced and transmitted from cell to cell. Endothelial cells are not passive but are rather active agents capable of reacting and changing the forces they experience. For instance, signaling vascular smooth muscles with vasoregulators to alter the local vascular tone leads to changes in hemodynamic conditions or by degrading and remodeling their extracellular matrix; thus, changing the mechanical and topographical properties of their basement membrane and the whole blood vessel. Ultimately, it is hoped through uncovering the mechanistic process by which endothelial cells sense their mechanical environment that targeted therapies can be developed to directly act on the mechanosensitive structures implicated in many vascular diseases. These therapies can be in the form of, pharmacological agents for interacting with cellular mechanosensitive structures, lifestyle changes such as exercise to disrupt hemodynamic conditions, or in biomedical devices designed to locally modify hemodynamic conditions.


Page 32 of 85

Page 33 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Responding to the Mechanical Microenvironment

Irrespective of the mechanism by which endothelial cells sense mechanical forces, these specialized cells respond to their mechanical microenvironment in specific ways. The influence of the individual hemodynamic forces (fluid flow, cyclic stretch, lateral pressure) and vessel forces (mechanical and topographical properties) from in vitro studies have all been reviewed extensively and their findings will be summarized. Less recognized are the interactive effects of these forces on endothelial cell behavior. The focus of this concluding section is to make prominent the nuanced responses of endothelial cells to combinations of different mechanical forces realized from recent in vitro investigations. The devices used for probing the effects of mechanical forces have been extensively reviewed in the literature and the interested reader is directed to such reviews for more details on device design, operation, and merits.225–228 In short, the effects of fluid flow have been investigated using several different devices including: parallel-plate flow chambers, cone and plate viscometers, parallel-plate viscometers, small-caliber tubes, and orbital shakers.228 Some of these devices have been designed to be high-throughput.229–232 The effects of stretch have been investigated using different mechanisms for straining a substrate; typically, this is achieved using a mechanical, pressure/pneumatic, or electromagnetic device to deform a membrane uni- or multiaxially.233 Both large-scale and microfluidic devices have been designed for studying cyclic stretch.88,233 The effects of hydrostatic pressure have been studied using specialized pressurized cell culture chambers, elevated cell culture media reservoirs, and hydroresistive elements.88,234,235 The effects of substrate stiffness has been studied using various hydrogels, polyurethanes, silicones, and engineered plastics.230,236–243 The common hydrogels employed have been polyacrylamides, methacrylated

polyethylene

glycols,

methacrylated

hyaluronic


acids,

and

poly(L-


lysine)/hyaluronan gels.230,237,238,242,244–246 The effects of topography have been investigated using engineered-topographies by soft lithographic methods and by electrospinning fibrous geometries247. The effects of multiple mechanical forces have been studied by combining the operating mechanisms of these individual devices.225,226

Fluid Flow

The endothelial cell response to fluid flow is unsurprisingly the most studied phenomenon due to its correlation with vascular disease and atherosclerosis formation.81,228,248 Chiu and Chien provide a comprehensive summary of the endothelial cell response to fluid flow from studies performed over the past 30 years and their analysis is summarized with minor supplementation from recent studies.228

Endothelial cells differentially respond to flow conditions. In general, under unidirectional laminar, high shear stress flow conditions, endothelial cells elongate, stiffen, and align in the direction of flow, express low rolling monocyte adhesion, low turnover (cell death), and reduced low-density lipoprotein permeability.228,249,250 Under these conditions, the expression of adhesion molecules and inflammatory and chemokine genes is low whereas the expression of antioxidant genes is high. Endothelialization is promoted, platelet aggregation inhibited, and heterogeneity among endothelial cells is low. Vascular smooth muscle cell activation is low and vasodilation is preferred.9,228 These characteristics of endothelial cells are principally considered to define the behavior of an atheroprotective phenotype.228 Additionally, unidirectional, steady laminar flow is recognized to induce directional remodeling of focal adhesions and stress fibers in the direction of


Page 34 of 85

Page 35 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


flow whereas no preferred distribution is observed for no flow and disturbed flow conditions.9,251 Most recently, Yoshino et al. investigated the effect of shear stress and shear stress gradients for laminar, high shear stress flow conditions. Surprisingly, the typical shear stresses that elicit elongation and alignment for uniform shear were unable to promote the same response in the presence of a shear stress gradient below a threshold shear stress. It was proposed that the shear stress gradient affects the endothelial cell strain field; thus, altering the localization of mechanosensing structures.252,253 In contrast, under disturbed or turbulent flow with low or reciprocating shear stress conditions, endothelial cells exhibit a polygonal shape and random orientation, express high rolling monocyte adhesion, high turnover, and elevated low-density lipoprotein permeability. Under these conditions, the expression of adhesion molecules, inflammatory, chemokine, and antioxidant genes is reversed. Endothelialization is retarded, platelet aggregation is promoted, and heterogeneity is high. Vascular smooth muscle cell activation is high and vasoconstriction is preferred.9,228 These characteristics of endothelial cells are principally considered to define the behavior of an atherogenic phenotype.228 Fluid flow induces a multitude of functional responses from endothelial cells, which have significant implications for vascular disease.

Cyclic Stretch

The endothelial cell response to cyclic stretch is as diverse and complex as that for fluid flow. The effects of cyclic stretch have been well-reviewed by several investigators and are summarized for the reader.9,11,101,153 Morphologically, endothelial cells elongate and align perpendicular to a uniaxial stretch direction and the response is enhanced with greater applied strain and frequency



as shown for strains up to 10% and frequencies up to 1.5 Hz.9,254–259 This has been visualized by actin stress fiber reorganization, which occurs in order to minimize tensile forces acting on the fibers9. Conversely, no alignment is achieved for non-uniaxial stretch.9 With this remodeling, the mechanical properties of endothelial cells change. For uniaxial cyclic stretch, endothelial cells stiffened and deformed less in response to pathological levels as compared to physiological levels of strain amplitude.260 Matrix remodeling is activated by stretch as observed by increased expression of matrix metalloproteinases (MMP); specifically, MMP-2 and MMP-14.111,261,262 Endothelial cell growth factors have also been found to activate in response to cyclic stretch; specifically, vascular endothelial growth factor and its receptor (VEGF and VEGF-R2), basic fibroblast growth factor (bFGF), angiopoietin-2, and platelet derived growth factor BB (PDGFBB).263–265 Activation of VEGF-R2 by pathological levels of cyclic stretch has been shown to mediate endothelial cell permeability.266 Vasodilators, endothelial nitric oxide synthase (eNOS) and prostacyclin (PGI2), and the vasoconstrictor, endothelin 1 (ET-1), all activate in response to stretch.267–271 Additionally, reactive oxygen species production was mediated by cyclic stretch being suppressed for physiological levels and enhanced by pathological levels.272,273 Recently, Pedrigi et al. showed that similar atherogenic phenotypes commonly associated with disturbed flow regimes can be induced by multidirectional stretch conditions.274 Lastly, endothelial cell proliferation has been observed to induce controlled proliferation for physiological levels and uncontrolled proliferation for pathological levels.275,276 Similarly, uniaxial cyclic stretch at physiological levels has been shown to protect endothelial cells from apoptosis while non-uniaxial and pathological levels of uniaxial cyclic stretch enhanced apoptosis.9,111 As with fluid flow, cyclic stretch elicits a range of functional responses from endothelial cells.


Page 36 of 85

Page 37 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Lateral Pressure

Quite possibly the least studied hemodynamic force is the effect of pressure on endothelial cell phenotype. Because of the similarities between endothelial cells exposed to pressure and the other hemodynamic forces, the endothelial response to pressure is suggestive of an active, proliferative phenotype.277 As with the effects of the other hemodynamic forces, the response to pressure is as variable. Endothelial cells cultured under sustained hydrostatic pressures of 147–19,998 Pa (1.1– 150 mmHg) for 1-9 days resulted in elongation with random orientation and reorganization of the cytoskeleton from a matrix of disperse fibers to arrays of aligned parallel stress fibers.278–280 Surface protein expression also has been investigated and hydrostatic pressure was shown to upregulate the expression of E-selectin and the integrin 5 subunit.277,281 Conversely, it was seen that VE-cadherin expression was downregulated in endothelial cells exposed to 6,666–19,998 Pa (50–150 mmHg) of pressure.282 Endothelial cell proliferation has been stimulated by sustained pressures of 147–25,497 Pa (1.1–191 mmHg) for 1-9 days and even caused the loss of contactinhibited growth of bovine endothelial cells, which was suggested to be due to the downregulation of VE-cadherin.234,277–279,282,283 Similarly, cyclic pressure modulated endothelial cell death and proliferation as a function of the mean pressure.284 This excessive proliferation has led to the observation of multilayered endothelial cell structures in cultures with pressure.282 This proliferative response is mediated by basic fibroblast growth factor (bFGF) and vascular endothelial growth factor C (VEGF-C).279,285,286 The effects of pressure appear to act on longer time scales (days) than those of shear or stretch (hours).277 However, it was shown that endothelin1 (ET-1) production increased with increasing pressure from 5,333–21,332 Pa (40–160 mmHg) within hours after exposure.287,288 Though in a dynamic experiment in which hydrostatic pressure



was ramped from 0 to 9333 (0 to 70 mmHg) and back down over 3 hours with each level being held for 1 hour, ET-1 production decreased at elevated pressures.289 Despite the well-known presence of pressure in the vasculature, in vitro studies of its effect on endothelial cells has been sparse.

Substrate stiffness

Many endothelial cell functions show some degree of substrate stiffness-dependence. From studies investigating blood vessel wall stiffening and cardiovascular disease, extracellular matrix stiffening has been shown to induce endothelial cell stiffening and increase endothelium permeability.236 Consequently, endothelial cell monolayer integrity negatively correlates with substrate stiffness.237 Recently, Lampi et al. investigated the effect of substrate stiffness heterogeneity using methacrylated hyaluronic acid hydrogels with elastic moduli of either 2.7 or 10.3 kPa patterned in a checkerboard arrangement.238 Increased stiffness heterogeneity resulted in the increased disruption of endothelial monolayer cell-cell junctions. Endothelial cells become stiffer when cultured on stiffer substrates in both a 2D and 3D setting as observed using collagen gels with elastic moduli of 1.7 and 9 kPa.239 Similarly, on polyacrylamide gels with elastic moduli of 0.1 – 10 kPa, endothelial cell stiffness increased with increasing substrate stiffness.230 Also in the same study, it was shown that cell spreading increased with substrate stiffness.230 Additionally, endothelial cell responsiveness to growth factors exhibits a substrate stiffness-dependence. Using polyacrylamide gels with elastic moduli of 4–125 kPa, endothelial cells differentially responded to vascular endothelial growth factor (VEGF) as a function of substrate stiffness.244 Endothelial cells were most responsive on soft gels as compared to the stiff gels. A similar stiffness-


Page 38 of 85

Page 39 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


dependence has been observed for hepatocyte growth factor.237 Endothelial cell migration is also mediated by substrate stiffness. In a modified cage assay, using polyacrylamide gels with elastic moduli ranging from 4–50 kPa endothelial cells differentially migrated as a function of substrate stiffness.245 Migration distance was greatest on the stiffest gels. Moreover, endothelial cell function is mediated by substrate stiffness. When cultured on poly(sodium p-styrene sulfonate) hydrogels with elastic modulus of 3–300 kPa, glycocalyx content increased with increasing gel stiffness as compared to tissue culture polystyrene.290 Additionally, both platelet adhesion and surface friction of endothelial cells was shown to depend on the substrate stiffness and expressed an optimal response for gels with an elastic modulus of ~60 kPa. Using poly(L-lysine)/hyaluronan gels with elastic moduli of 200–430 kPa, nitric oxide production negatively correlated with substrate stiffness.237 Endothelial cell proliferation is also mediated by substrate stiffness. Using polyacrylamide gels with elastic moduli of 1.72–21.5 kPa and glass slides with an elastic modulus of >60 GPa, endothelial cell proliferation increased with increasing substrate stiffness. However, no statistical difference in proliferation was found between the stiffest gel and glass indicating that the plateau for stiffness-mediated proliferation may have already been reached.291 Similarly, in another study using poly(L-lysine)/hyaluronan gels with elastic moduli of 200–430 kPa, the same stiffness-mediated proliferation trend was observed.237 However, this was shown for much stiffer gels beyond the plateauing threshold observed with polyacrylamide gels, which may be the result of several potential differences such as substrate chemistry. Substrate stiffness seems to regulate all facets of endothelial cell function from monolayer integrity to glycocalyx layer composition.



Topography

Endothelial cell response to topographic features is similarly diverse but is seemingly less defined. Greiner et al. reviewed the effect of engineered topographic features on endothelial cells.247 The engineered topographies investigated included; gold nano-patterned dots, nanowires, micropillars, microposts, microgrooves, and nanofibers.247 Proliferation has been observed to depend on both the shape and size of the topography and in general increases on nanotopographic features.247 Similarly, adhesion has been seen to depend on the structure and size of the surface features.247 A commonly employed engineered topography has been the use of well-defined grooves or channels. Grooved topographies directed endothelial cell alignment and elongation along the groove direction.247 Endothelial monolayer antithrombogenicity was also enhanced on grooves and showed a feature size dependence.292 Endothelial cell motility depended on the topographic features and migration was directed by aligned topographies such as grooves.247 The morphological changes were typified by alignment and elongation of actin stress fibers in the groove direction.247 Recently, Lamichhane et al. investigated the topographic influence of polytetrafluoroethylene (PTFE) on endothelial cells and found increased proliferation on electrospun PTFE over both expanded and flat PTFE.293 Conversely, topographies mimicking basement membrane features in NOA81, a rigid mercapto-ester polymer, reduced proliferation rate and expression of matrix metalloproteinase-2, enhanced migration rate, and did not effect PECAM-1 expression.294 Interestingly, Cutiongco et al. investigated the effects of hierarchical grooved and microlens curvature on healthy and diabetic endothelial cells.295 It was shown that hierarchical grooves were necessary to elicit consistent effects in diabetic endothelial cells observed as reduced uptake of oxidized low-density lipoprotein, decreased immune activation, and


Page 40 of 85

Page 41 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


accelerated migration. Concave microlens were shown to deteriorate diabetic endothelial cell function. Endothelial cells exhibit contact guided functional responses to both native and engineered topographies.

Combinations

The combined effects of fluid flow and cyclic stretch influence both endothelial cell morphology and vasoregulation. Synergistic effects for both alignment and elongation were observed after exposure to physiological levels of steady flow and cyclic stretch.296,297 Owatverot et al. investigated the relative strengths of the two stimuli in reinforcing and counteracting combinations.298 Equipotent levels, stimuli of different type able to elicit equivalent responses, of each stimulus were 8 Pa (80 dynes/cm2) shear stress for steady flow, 2 Pa/s (20 dynes/cm2-s) shear stress for oscillating flow, and 2% strain at 1 Hz for cyclic stretch. The reinforcing combination of equipotent steady flow and cyclic stretch did not produce an additive response for alignment and elongation. In contrast, the reinforcing combination of equipotent oscillating flow and cyclic stretch did show an additive response in both alignment and elongation. The response to the counteracting equipotent combinations was essentially canceled resulting in random alignment and morphology. For combinations of steady flow and anisotropic biaxial strains, alignment showed a strain dependence at very low shear stress while the alignment response transitioned to a shear stress dependence when exposed to low shear stress.299 Vasoregulation was shown to depend on the stress phase angle (SPA) between oscillatory flow with 1 Pa (10 dynes/cm2) and 8% cyclic stretch at 1 Hz.300 Prostacyclin production rate drastically decreased for increasingly negative SPA as compared to a steady flow control. Nitric oxide production rate decreased and trended slightly



with increasingly negative SPA as compared to steady flow. ET-1 production rate increased with increasingly negative SPA as compared to steady flow. Across different studies, trends for endothelial nitric oxide synthase and ET-1 expression agreed; despite, differences in shear stress magnitude, strain magnitude, and SPA.267,301 In a later study, it was found that a SPA of -180 elicited an atherogenic phenotype in bovine aortic endothelial cells.302 Static and temporal combinations of fluid flow and cyclic stretch have demonstrated unique responses deviating from those of each individual stimulus.

Studies on the effect of fluid flow and hydrostatic pressure are sparse. Human umbilical vein endothelial cells exposed to steady flow with 1.208 Pa (12.08 dynes/cm2) shear stress and hydrostatic pressure ranging from 5560–14972 Pa (41.7–112.3 mmHg) exhibited reduced VEcadherin expression with increased hydrostatic pressure.235 Alignment and elongation both decreased upon stimulation of flow with only a weak dependence on hydrostatic pressure235. Much has yet to be learned about the combinatorial effects of fluid flow and hydrostatic pressure on endothelial cells.

The addition of surface topography to endothelial cells exposed to fluid flow demonstrated both synergistic and antagonistic effects depending on the relative orientation of the topographies and their degree of anisotropy. This was first demonstrated with endothelial cells by studying combinations of grooved topographies and steady flow with 1.35–5.8 Pa (13.5–58 dynes/cm2) shear stress.303 It was shown that topographies can either add or compete with the effects of fluid flow. Morgan et al. later examined combinations of grooved or pitted topographies and steady flow with 2.0 Pa (20 dynes/cm2) shear stress.304 The authors reported that the combined effects were


Page 42 of 85

Page 43 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


both feature- and size-dependent (Figure 6). Isotropic features (pits) combined with shear did not influence endothelial cell alignment while the anisotropic features (grooves) with fluid flow did influence alignment. Further, with reduced groove size (800 nm), fluid flow dominated endothelial cell alignment while with enlarged groove size (2000 nm), topography dominated endothelial cell alignment.304 Moreover, a synergistic response was seen for endothelial cell alignment when the substrate grooves were parallel to the flow direction and antagonistic when the grooves were perpendicular to the flow direction. Within this comprehensive study, endothelial cell motility was also investigated.304 Migration velocity was unchanged for grooves perpendicular to flow for all groove sizes as compared to a flat surface. Though, migration direction and path tortuosity both increased for larger groove sizes, which indicates the competing effects of anisotropic topography with fluid flow. The impact on migration and monolayer integrity was further investigated with grooved topographies and steady flow with 1.4 Pa (14 dynes/cm2) shear stress.305 Migratory action was assessed by a wound healing/scratch assay, which showed improved migration (healing) by the presence of the grooved topography parallel to the flow direction.305 The presence of both flow and topography promoted collective monolayer migration over individual cell migration. This effect was further reinforced by investigating the expression of VE-cadherin, which was shown to have an optimal level regulated by enhancement from the topographic cues and diminishment from the flow.305 The extent of this effect was later investigated with grooved topographies and steady flow with 1.4–10 Pa (14–100 dynes/cm2) shear stress.306 Monolayer integrity, the intactness of the cultured endothelial monolayer, was enhanced on grooves parallel to the flow direction up to 8 Pa (80 dynes/cm2) shear stress. Likewise, cell density was shown to remain stable up to the maximum tested shear stress of 10 Pa (100 dynes/cm2). As previously described, endothelial cells cultured on grooves parallel to the flow direction at low shear stress showed enhanced alignment; however,



at shear stresses between 4 – 8 Pa (40–80 dynes/cm2) alignment was entirely lost and regained at shear stresses of 10 Pa (100 dynes/cm2). Surprisingly, at a shear stresses greater than or equal to 4 Pa (40 dynes/cm2) a collection of endothelial cells aligned perpendicular to the flow direction and parallel to the groove direction.306 Pre-culturing an endothelial layer on the grooved substrate and then subjecting it to flow perpendicular to the groove direction yielded sustained monolayer integrity.306 The impact of geometry was studied using wavy-grooved and square-grooved topographies and steady fluid flow with 1–10 Pa (10–100 dynes/cm2) shear stress.307 Improved monolayer integrity was shown for the wavy-grooved topography oriented both parallel and perpendicular to the flow direction at all levels of shear stress as compared to the square-grooved topography and flat substrate. The translation of these effects in engineered-topographies to more physiological topographies was further investigated with electrospun scaffolds of variable fiber alignment and steady fluid flow with 2 or 4 Pa (20 or 40 dynes/cm2) shear stress.308 Trends in alignment, elongation, monolayer integrity, and VE-cadherin expression largely mirrored those for grooved topographies. Surface topographies can be designed to compete with the effects of fluid flow.


Page 44 of 85

Page 45 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 6. Endothelial cells respond both synergistically and antagonistically in response substrate topography and fluid flow. Ridges (grooves) oriented parallel to the direction of flow demonstrated a synergistic effect, whereas orientations perpendicular to flow had an antagonistic effect. White arrows indicate the direction of the ridges (grooves), the direction of flow is indicated with a black arrow positioned under the fluorescence microscopy images. Cellular alignment changes were demonstrated by reorientation of actin stress fibers (red), microtubules (green), and nuclear morphology (blue). Scale Bar = 50 μm. Reproduced with permission from ref 304. Copyright 2012 Elsevier.



Substrate stiffness can work in tandem with fluid flow to moderate morphology, vasoregulation, and inflammation. Confluent monolayers aligned in the direction of flow irrespective of substrate stiffness for cultures on polyacrylamide hydrogels with elastic modulus of 2.5 or 10 kPa and steady fluid flow with 1.2 Pa (12 dynes/cm2) shear stress.309 Membrane perimeter, elongation, barrier integrity, and nitric oxide production were all enhanced on more compliant substrates while under flow conditions. Conversely, alignment of subconfluent cells showed a non-linear dependence on both shear stress and substrate stiffness for cultures on polyacrylamide hydrogels with elastic modulus of 0.1, 2.5, or 10 kPa and steady fluid flow with 0.6, 1.2, 1.8, and 2.2 Pa (6, 12, 18, and 22 dynes/cm2) shear stresses.230 As the substrate increased in compliance, the magnitude of shear stress necessary to achieve alignment with the flow direction increased (Figure 7). In a similar fashion, both cell area and cell stiffness showed a non-linear dependence on both shear stress and substrate stiffness. Moreover, substrate stiffness had the ability to reduce endothelial cell sensitivity to tumor necrosis factor alpha (TNF-), an inflammation cytokine. Substrate stiffness can moderate the effects of fluid flow.


Page 46 of 85

Page 47 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 7. Combinations of substrate stiffness and fluid flow influence endothelial cell morphology in a non-linear manner. Endothelial cells were stained with phalloidin (red) for F-actin fibers and DAPI (blue) for the nuclear DNA (each image is 75 × 75 μm). Adapted from ref 230 with permission of The Royal Society of Chemistry.

Understanding of the combined effects of cyclic stretch and substrate stiffness on endothelial cell behavior is minimal; despite, a significant interaction being observed. The combination of cyclic stretch and substrate stiffness coordinate to protect endothelial cell monolayers against inflammation agonists.310 Membrane integrity was enhanced on softer substrates following thrombin-induced gap formation for monolayers cultured on polyacrylamide hydrogels with elastic modulus of 5 and 15 kPa and 5% biaxial cyclic stretch at 0.25 Hz. Much has yet to be



learned about the combinatorial effects of cyclic stretch and substrate stiffness on endothelial cell behavior.

Substrate properties interact to elicit unique responses from endothelial cells in the absence of hemodynamic forces. The impact of these combined stimuli on endothelial cell inflammation has largely been investigated. Inflammatory and angiogenic cytokine and chemokine production was mediated by surface topography and slightly by substrate stiffness for cultures on poly(urethane acrylates) with supraphysiological stiffness and grooved topographies with varying groove spacing.240 The effects attributed to surface topography disappeared upon exposure to TNF-. Similar results were observed for endothelial cell alignment on polydimethylsiloxane elastomers with grooves of varying geometries and with varying elastic modulus (300–2300 kPa).241 Endothelial cell alignment showed minimal dependence on substrate stiffness and was largely dependent on the groove width (5–20 µm) and height (1.5 and 5 µm). Ding et al. conducted an expansive investigation on the interactive effects of substrate topography, stiffness, and matrix composition using a high-throughput method for printing specific combinations of matrix proteins on poly(ethylene glycol) dimethacrylate hydrogels.242 Three different hydrogels were prepared, a soft, smooth gel with elastic modulus of 1.5 kPa, a stiff, smooth gel with elastic modulus of 15 kPa, and a soft electrospun fibrous gel with elastic modulus of 1.5 kPa. To assess the role of the extracellular matrix composition, the different gels were templated with 21 paired combinations of gelatin (used as a control), collagen type I, collagen type III, collagen type IV, laminin, and fibronectin. The degree of cell attachment was highly dependent on both surface topography and composition, but not on the range of substrate stiffness that was included in the investigation. Proliferation showed a similar matrix compositional dependence for both soft and stiff gels with


Page 48 of 85

Page 49 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


stiffer gels eliciting a greater response. A fibrous topography reduced proliferation for all matrix compositions. Nuclear factor kappa B (NF- B), an indicator of inflammation, showed a dependence on both matrix composition and stiffness and not on surface topography. Response to TNF- was mediated by all three factors. It should be noted that these cultures were all contained in the same vessel being exposed to the same cell culture media. Surface topography and substrate stiffness in combination mediate endothelial cell response.

Several studies have developed bioreactors capable of producing complex pulsatile flow, pulsatile pressure, and cyclic stretch profiles; however, their utility beyond proof-of-concept investigations has been limited.311–315 These systems can provide a realistic complex mechanical microenvironment; though, decoupling of the three hemodynamic forces is not without its challenges. The benefit of these in vitro systems is that they can confer complex mechanical forces to cultured endothelial cells without the confounding effects of other factors (chemical, physiological, and biological) acting on the endothelium in an in vivo study. The proof-of-concept studies presented here reiterated that endothelial cells showed an alignment dependence with the applied forces. Exposure to all pulsatile flow, pulsatile pressure, and cyclic stretch demonstrated that the combination of forces differentially altered vasoregulatory and fibrinolytic functions of endothelial cells.316,317 Much is left to be revealed about the interplay between mechanical forces and their combined effects on endothelial cell function.



Conclusions

In 2009 in the textbook Cellular Mechanotransduction, Tarbell expressed the confusing behavior of the field at the end of his chapter, “The Interaction between Fluid-Wall Shear Stress and Solid Circumferential Strain Affects Endothelial Cell Mechanobiology”,

There appear to be at least two reasons for the lack of emphasis on combined forces in the literature. One is related to the difficulty of applying combined forces experimentally in vitro. Further emphasis on the development of convenient systems for application of simultaneous forces in vitro will surely advance the field. The second is simply a lack of appreciation of the possibility for unique endothelial cell responses to combined forces, not characteristic of either force alone.318

Since his statement, there have been several devices developed capable of investigating pairs of these mechanical forces, which have revealed many unique responses.225,227,315,319,230,242,300,304,311– 314

Nonetheless, it seems these devices are designed, validated, and used for a singular study. The

second challenge of appreciation is slowly being overcome. Largely the data on the effects of combined mechanical forces on endothelial cells comes from a handful of studies conducted in the last few years, primarily focused on the interaction between fluid flow and cyclic stretch. Few probed the other pairwise combinations of which several to the author’s knowledge have yet to be investigated in endothelial cells; namely, pressure with stiffness, topography, or stretch, and stretch with topography. Though some have been studied for other cell types.226 Two additional challenges that need to be a considered are 1) the diversity in endothelial cell type and 2) the large number of experimental conditions required for testing combinations of mechanical forces. Endothelial cells


Page 50 of 85

Page 51 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


display tremendous heterogeneity based on their location in the body.320 Meanwhile, studies almost exclusively use human umbilical vein endothelial cells, human aortic endothelial cells, or bovine aortic endothelial cells. These cell types are likely most commonly used because of their ease for growth in culture. Because of the spatial diversity in the mechanical microenvironment and the epigenetic changes encoded in primary cells, the field needs to pair specific endothelial cell type with their physiologically-relevant mechanical forces to ensure the greatest translatability of these in vitro studies to location-specific behavior.321 The large number of experimental conditions can be accounted for by using experimental design methodologies in conjunction with high-throughput devices. Together these methods can reduce the number of needed experiments, account for interactions between forces, and reduce the time of experimentation.242,322 This review considered the effects of arterial and venous mechanical forces and not those experienced in capillaries (microvasculature). This separate environment exposes endothelial cells to different cellular interactions and mechanics, namely a 3D matrix during angiogenesis. The interested reader is directed to a collection of numerous reviews and studies for further information on microvasculature mechanical forces and their effects on endothelial cell response and angiogenesis.323–335 The vascular mechanical microenvironment intrinsically exposes endothelial cells to a complex milieu of forces, which are detected using an assortment of mechanosensing structures, and are transduced into functional responses. Targeted therapies can be designed to influence endothelial cell mechanosensors to mitigate the effects of pathological mechanical forces.

Biomaterials Perspective Current, vascular implants face issues with long-term patency potentially because the complexity of the mechanical microenvironment is overly simplified or reduced during their design.336,337 For



example, synthetic vascular implants largely fail to mimic the anisotropy, non-linearity, and viscoelasticity of the native blood vessel, which all contribute to their issues with long-term patency.338 Blood vessel extracellular matrix proteins are important regulators of endothelial cell function.17 Identifying the proteins that adsorb to the surface of the biomaterial and that are secreted by the cells in response to the biomaterial is necessary. Endothelial cell functionality relies on the complexity of mechanics, topography, and chemistry.242 Recently, it was shown that monolayers of cells can detect substrate thicknesses of hundreds of microns.200 The endothelial cell basement membrane is only a few hundred nanometers, implying that the mechanical properties of the entire vessel are likely paramount because the cells would be capable of sensing the mechanical properties of the outer layers of the blood vessel.141,339 In a similar manner, even when mechanical properties can largely be mimicked in tissue-engineered vascular grafts, undesirable hemodynamic forces acting in the grafts can impede their performance.340 Because of the interactive effects between substrate stiffness and fluid flow, conceivably future vascular implants could be designed in which the mechanical properties of a soft biomaterial could mediate the effects of the hemodynamic conditions. Expanding on this further, the topographic features on the surface of a vascular implant could guide endothelial cells for improved endothelialization and mitigate the adverse effects of atherogenic low shear stress, disturbed flows. Moreover, combinations of both topographic features and mechanical properties could be used to reconcile the effects of adverse hemodynamic forces. Having a better understanding of the interactions between controllable vessel wall forces (substrate stiffness, surface topography) and the largely uncontrollable hemodynamic forces (fluid flow, cyclic stretch, lateral pressure), vascular biomaterials and biomedical devices could be engineered from the bottom-up to account for the endothelial cellular response.


Page 52 of 85

Page 53 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


References (1)

Huang, G.; Wang, L.; Wang, S.; Han, Y.; Wu, J.; Zhang, Q.; Xu, F.; Lu, T. J. Engineering ThreeDimensional Cell Mechanical Microenvironment with Hydrogels. Biofabrication 2012, 4 (4). DOI:10.1088/1758-5082/4/4/042001.

(2)

Nagelkerke, A.; Bussink, J.; Rowan, A. E.; Span, P. N. The Mechanical Microenvironment in Cancer: How Physics Affects Tumours. Semin. Cancer Biol. 2015, 35, 62–70. DOI:10.1016/j.semcancer.2015.09.001.

(3)

Discher, D. E. Tissue Cells Feel and Respond to the Stiffness of Their Substrate. Science (80-. ). 2005, 310 (5751), 1139–1143. DOI:10.1126/science.1116995.

(4)

Paluch, E. K.; Nelson, C. M.; Biais, N.; Fabry, B.; Moeller, J.; Pruitt, B. L.; Wollnik, C.; Kudryasheva, G.; Rehfeldt, F.; Federle, W. Mechanotransduction: Use the Force(S). BMC Biol. 2015, 13 (1), 1–14. DOI:10.1186/s12915-015-0150-4.

(5)

Engler, A. J.; Sen, S.; Sweeney, H. L.; Discher, D. E. Matrix Elasticity Directs Stem Cell Lineage Specification. Cell 2006, 126 (4), 677–689. DOI:10.1016/j.cell.2006.06.044.

(6)

Levental, I.; Georges, P. C.; Janmey, P. A. Soft Biological Materials and Their Impact on Cell Function. Soft Matter 2007, 3 (3), 299–306. DOI:10.1039/B610522J.

(7)

Chaudhuri, O.; Gu, L.; Klumpers, D.; Darnell, M.; Bencherif, S. A.; Weaver, J. C.; Huebsch, N.; Lee, H. P.; Lippens, E.; Duda, G. N.; et al. Hydrogels with Tunable Stress Relaxation Regulate Stem Cell Fate and Activity. Nat. Mater. 2016, 15 (3), 326–334. DOI:10.1038/nmat4489.

(8)

Yang, C.; Tibbitt, M. W.; Basta, L.; Anseth, K. S. Mechanical Memory and Dosing Influence Stem Cell Fate. Nat. Mater. 2014, 13 (6), 645–652. DOI:10.1038/nmat3889.

(9)

Chien, S. Mechanotransduction and Endothelial Cell Homeostasis: The Wisdom of the Cell. Am. J. Physiol. Circ. Physiol. 2007, 292 (3), H1209–H1224. DOI:10.1152/ajpheart.01047.2006.

(10)

Gimbrone, M. A.; García-Cardeña, G. Endothelial Cell Dysfunction and the Pathobiology of Atherosclerosis. Circ. Res. 2016, 118 (4), 620–636. DOI:10.1161/CIRCRESAHA.115.306301.

(11)

Jufri, N. F.; Mohamedali, A.; Avolio, A.; Baker, M. S. Mechanical Stretch: Physiological and Pathological Implications for Human Vascular Endothelial Cells. Vasc. Cell 2015, 7 (1), 8. DOI:10.1186/s13221-0150033-z.

(12)

Demer, L. L.; Tintut, Y. Vascular Calcification: Pathobiology of a Multifaceted Disease. Circulation 2008,



117 (22), 2938–2948. DOI:10.1161/CIRCULATIONAHA.107.743161. (13)

Akyildiz, A. C.; Speelman, L.; Gijsen, F. J. H. Mechanical Properties of Human Atherosclerotic Intima Tissue. J. Biomech. 2014, 47 (4), 773–783. DOI:10.1016/j.jbiomech.2014.01.019.

(14)

Ponticos, M.; Smith, B. D. Extracellular Matrix Synthesis in Vascular Disease: Hypertension, and Atherosclerosis. J. Biomed. Res. 2014, 28 (1), 25–39. DOI:10.7555/JBR.27.20130064.

(15)

Chistiakov, D. A.; Sobenin, I. A.; Orekhov, A. N. Vascular Extracellular Matrix in Atherosclerosis. Cardiol. Rev. 2013, 21 (6), 270–288. DOI:10.1097/CRD.0b013e31828c5ced.

(16)

Yurdagul, A.; Finney, A. C.; Woolard, M. D.; Orr, A. W. The Arterial Microenvironment: The Where and Why of Atherosclerosis. Biochem. J. 2016, 473 (10), 1281–1295. DOI:10.1042/BJ20150844.

(17)

Eble, J.; Niland, S. The Extracellular Matrix of Blood Vessels. Curr. Pharm. Des. 2009, 15 (12), 1385– 1400. DOI:10.2174/138161209787846757.

(18)

Tarbell, J. M.; Ebong, E. E. Endothelial Glycocalyx Structure and Role in Mechanotransduction. In Hemodynamics and Mechanobiology of Endothelium; WORLD SCIENTIFIC: Singapore, 2010; pp 69–95. DOI:10.1142/9789814280426_0003.

(19)

Tarbell, J. M.; Simon, S. I.; Curry, F.-R. E. Mechanosensing at the Vascular Interface. Annu. Rev. Biomed. Eng. 2014, 16 (1), 505–532. DOI:10.1146/annurev-bioeng-071813-104908.

(20)

Reitsma, S.; Slaaf, D. W.; Vink, H.; van Zandvoort, M. A. M. J.; oude Egbrink, M. G. A. The Endothelial Glycocalyx: Composition, Functions, and Visualization. Pflügers Arch. - Eur. J. Physiol. 2007, 454 (3), 345–359. DOI:10.1007/s00424-007-0212-8.

(21)

Schött, U.; Solomon, C.; Fries, D.; Bentzer, P. The Endothelial Glycocalyx and Its Disruption, Protection and Regeneration: A Narrative Review. Scand. J. Trauma. Resusc. Emerg. Med. 2016, 24 (1), 48. DOI:10.1186/s13049-016-0239-y.

(22)

Weinbaum, S.; Tarbell, J. M.; Damiano, E. R. The Structure and Function of the Endothelial Glycocalyx Layer. Annu. Rev. Biomed. Eng. 2007, 9 (1), 121–167. DOI:10.1146/annurev.bioeng.9.060906.151959.

(23)

Castellot, J. J.; Rosenberg, R. D.; Karnovsky, M. J. Endothelium, Heparin, and the Regulation of Vascular Smooth Muscle Cell Growth. In Biology of Endothelial Cells; Jaffe, E. A., Ed.; Developments in Cardiovascular Medicine; Springer US: Boston, MA, 1984; Vol. 27, pp 118–128. DOI:10.1007/978-1-46132825-4.


Page 54 of 85

Page 55 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(24)

Bazzoni, G. Endothelial Cell-to-Cell Junctions: Molecular Organization and Role in Vascular Homeostasis. Physiol. Rev. 2004, 84 (3), 869–901. DOI:10.1152/physrev.00035.2003.

(25)

Mundi, S.; Massaro, M.; Scoditti, E.; Carluccio, M. A.; van Hinsbergh, V. W. M.; Iruela-Arispe, M. L.; De Caterina, R. Endothelial Permeability, LDL Deposition, and Cardiovascular Risk Factors—a Review. Cardiovasc. Res. 2018, 114 (1), 35–52. DOI:10.1093/cvr/cvx226.

(26)

Wallez, Y.; Huber, P. Endothelial Adherens and Tight Junctions in Vascular Homeostasis, Inflammation and Angiogenesis. Biochim. Biophys. Acta - Biomembr. 2008, 1778 (3), 794–809. DOI:10.1016/j.bbamem.2007.09.003.

(27)

Mundi, S.; Massaro, M.; Scoditti, E.; Carluccio, M. A.; van Hinsbergh, V. W. M.; Iruela-Arispe, M. L.; De Caterina, R. Endothelial Permeability, LDL Deposition, and Cardiovascular Risk Factors—a Review. Cardiovasc. Res. 2018, 114 (1), 35–52. DOI:10.1093/cvr/cvx226.

(28)

Mestas, J.; Ley, K. Monocyte-Endothelial Cell Interactions in the Development of Atherosclerosis. Trends Cardiovasc. Med. 2008, 18 (6), 228–232. DOI:10.1016/j.tcm.2008.11.004.

(29)

Moore, K. J.; Tabas, I. Macrophages in the Pathogenesis of Atherosclerosis. Cell 2011, 145 (3), 341–355. DOI:10.1016/j.cell.2011.04.005.

(30)

Mestas, J.; Ley, K. Monocyte-Endothelial Cell Interactions in the Development of Atherosclerosis. Trends Cardiovasc. Med. 2008, 18 (6), 228–232. DOI:10.1016/j.tcm.2008.11.004.

(31)

Moore, K. J.; Tabas, I. Macrophages in the Pathogenesis of Atherosclerosis. Cell 2011, 145 (3), 341–355. DOI:10.1016/j.cell.2011.04.005.

(32)

Gimbrone, M. A.; García-Cardeña, G. Endothelial Cell Dysfunction and the Pathobiology of Atherosclerosis. Circ. Res. 2016, 118 (4), 620–636. DOI:10.1161/CIRCRESAHA.115.306301.

(33)

Xu, J.; Shi, G.-P. Vascular Wall Extracellular Matrix Proteins and Vascular Diseases. Biochim. Biophys. Acta - Mol. Basis Dis. 2014, 1842 (11), 2106–2119. DOI:10.1016/j.bbadis.2014.07.008.

(34)

Aumailley, M.; Smyth, N. The Role of Laminins in Basement Membrane Function. J. Anat. 1998, 193 (1), 1–21. DOI:10.1017/S0021878298003720.

(35)

Smyth, N.; Vatansever, H. S.; Murray, P.; Meyer, M.; Frie, C.; Paulsson, M.; Edgar, D. Absence of Basement Membranes after Targeting the LAMC1 Gene Results in Embryonic Lethality Due to Failure of Endoderm Differentiation. J. Cell Biol. 1999, 144 (1), 151–160. DOI:10.1083/jcb.144.1.151.



(36)

Pulkkinen, L.; Uitto, J. Mutation Analysis and Molecular Genetics of Epidermolysis Bullosa. Matrix Biol. 1999, 18 (1), 29–42. DOI:10.1016/S0945-053X(98)00005-5.

(37)

Kalluri, R. Basement Membranes: Structure, Assembly and Role in Tumour Angiogenesis. Nat. Rev. Cancer 2003, 3 (6), 422–433. DOI:10.1038/nrc1094.

(38)

Belkin, A. M.; Stepp, M. A. Integrins as Receptors for Laminins. Microsc. Res. Tech. 2000, 51 (3), 280– 301. DOI:10.1002/1097-0029(20001101)51:33.0.CO;2-O.

(39)

Beauvais, D. M.; Rapraeger, A. C. Syndecans in Tumor Cell Adhesion and Signaling. Reprod. Biol. Endocrinol. 2004, 2 (1), 3. DOI:10.1186/1477-7827-2-3.

(40)

Aumailley, M.; Battaglia, C.; Mayer, U.; Reinhardt, D.; Nischt, R.; Timpl, R.; Fox, J. W. Nidogen Mediates the Formation of Ternary Complexes of Basement Membrane Components. Kidney Int. 1993, 43 (1), 7–12. DOI:10.1038/ki.1993.3.

(41)

Ho, M. S. P.; Böse, K.; Mokkapati, S.; Nischt, R.; Smyth, N. Nidogens-Extracellular Matrix Linker Molecules. Microsc. Res. Tech. 2008, 71 (5), 387–395. DOI:10.1002/jemt.20567.

(42)

Marneros, A. G. Physiological Role of Collagen XVIII and Endostatin. FASEB J. 2005, 19 (7), 716–728. DOI:10.1096/fj.04-2134rev.

(43)

Segev, A.; Nili, N.; Strauss, B. H. The Role of Perlecan in Arterial Injury and Angiogenesis. Cardiovasc. Res. 2004, 63 (4), 603–610. DOI:10.1016/j.cardiores.2004.03.028.

(44)

Knox, S. M.; Whitelock, J. M. Perlecan: How Does One Molecule Do so Many Things? Cell. Mol. Life Sci. 2006, 63 (21), 2435–2445. DOI:10.1007/s00018-006-6162-z.

(45)

Farach-Carson, M. C.; Carson, D. D. Perlecan - A Multifunctional Extracellular Proteoglycan Scaffold. Glycobiology 2007, 17 (9), 897–905. DOI:10.1093/glycob/cwm043.

(46)

Bix, G.; Iozzo, R. V. Novel Interactions of Perlecan: Unraveling Perlecan’s Role in Angiogenesis. Microsc. Res. Tech. 2008, 71 (5), 339–348. DOI:10.1002/jemt.20562.

(47)

Zhou, Z.; Wang, J.; Cao, R.; Morita, H.; Soininen, R.; Chan, K. M.; Liu, B.; Cao, Y.; Tryggvason, K. Impaired Angiogenesis, Delayed Wound Healing and Retarded Tumor Growth in Perlecan Heparan SulfateDeficient Mice. Cancer Res. 2004, 64 (14), 4699–4702. DOI:10.1158/0008-5472.CAN-04-0810.

(48)

Blann, A. D. Plasma von Willebrand Factor, Thrombosis, and the Endothelium: The First 30 Years. Thromb. Haemost. 2005, 95 (4), 715–719. DOI:10.1160/TH05-07-0527.


Page 56 of 85

Page 57 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(49)

Franchini, M.; Lippi, G. Von Willebrand Factor and Thrombosis. Ann. Hematol. 2006, 85 (7), 415–423. DOI:10.1007/s00277-006-0085-5.

(50)

Kurkinen, M.; Vaheri, A.; Roberts, P. J.; Stenman, S. Sequential Appearance of Fibronectin and Collagen in Experimental Granulation Tissue. Lab. Invest. 1980, 43 (1), 47–51.

(51)

Labat-Robert, J.; Szendroi, M.; Godeau, G.; Robert, L. Comparative Distribution Patterns of Type I and III Collagens and Fibronectin in Human Arteriosclerotic Aorta. Pathol. Biol. (Paris). 1985, 33 (4), 261–265.

(52)

Murata, K.; Motayama, T.; Kotake, C. Collagen Types in Various Layers of the Human Aorta and Their Changes with the Atherosclerotic Process. Atherosclerosis 1986, 60 (3), 251–262.

(53)

Shekhonin, B. V.; Domogatsky, S. P.; Idelson, G. L.; Koteliansky, V. E.; Rukosuev, V. S. Relative Distribution of Fibronectin and Type I, III, IV, V Collagens in Normal and Atherosclerotic Intima of Human Arteries. Atherosclerosis 1987, 67 (1), 9–16. DOI:10.1016/0021-9150(87)90259-0.

(54)

Aoki, T.; Yamamoto, K. Fundamentals of Physiology and Biology of Vascular System. In Vascular Engineering; Tanishita, K., Yamamoto, K., Eds.; Springer Japan: Tokyo, 2016; pp 47–68. DOI:10.1007/978-4-431-54801-0.

(55)

Vanhoutte, P. M.; Zhao, Y.; Xu, A.; Leung, S. W. S. Thirty Years of Saying NO. Circ. Res. 2016, 119 (2), 375–396. DOI:10.1161/CIRCRESAHA.116.306531.

(56)

Furchgott, R. F.; Vanhoutte, P. M. Endothelium-Derived Relaxing and Contracting Factors. FASEB J. 1989, 3 (9), 2007–2018. DOI:10.1096/fasebj.3.9.2545495.

(57)

Vanhoutte, P. M. How We Learned to Say NO. Arterioscler. Thromb. Vasc. Biol. 2009, 29 (8), 1156–1160. DOI:10.1161/ATVBAHA.109.190215.

(58)

Moncada, S. Nitric Oxide in the Vasculature: Physiology and Pathophysiology. Ann. N. Y. Acad. Sci. 1997, 811, 60–69.

(59)

Murad, F. Discovery of Some of the Biological Effects of Nitric Oxide and Its Role in Cell Signaling (Nobel Lecture). Angew. Chemie Int. Ed. 1999, 38 (13–14), 1856–1868. DOI:10.1002/(SICI)15213773(19990712)38:13/143.0.CO;2-D.

(60)

Loscalzo, J. The Identification of Nitric Oxide as Endothelium-Derived Relaxing Factor. Circ. Res. 2013, 113 (2), 100–103. DOI:10.1161/CIRCRESAHA.113.301577.

(61)

Ignarro, L. J. Biosynthesis and Metabolism of Endothelium-Derived Nitric Oxide. Annu. Rev. Pharmacol.



Toxicol. 1990, 30 (1), 535–560. DOI:10.1146/annurev.pa.30.040190.002535. (62)

Thyberg, J.; Blomgren, K.; Roy, J.; Tran, P. K.; Hedin, U. Phenotypic Modulation of Smooth Muscle Cells after Arterial Injury Is Associated with Changes in the Distribution of Laminin and Fibronectin. J. Histochem. Cytochem. 1997, 45 (6), 837–846. DOI:10.1177/002215549704500608.

(63)

Moiseeva, E. Adhesion Receptors of Vascular Smooth Muscle Cells and Their Functions. Cardiovasc. Res. 2001, 52 (3), 372–386. DOI:10.1016/S0008-6363(01)00399-6.

(64)

Campbell, J. H.; Campbell, G. R. Endothelial Cell Influences on Vascular Smooth Muscle Phenotype. Annu. Rev. Physiol. 1986, 48 (1), 295–306. DOI:10.1146/annurev.ph.48.030186.001455.

(65)

Mithieux, S. M.; Weiss, A. S. Elastin. Adv. Protein Chem. 2005, 70, 437–461. DOI:10.1016/S00653233(05)70013-9.

(66)

Brooke, B. Extracellular Matrix in Vascular Morphogenesis and Disease: Structure versus Signal. Trends Cell Biol. 2003, 13 (1), 51–56. DOI:10.1016/S0962-8924(02)00007-7.

(67)

Ito, S. Inhibitory Effect of Type 1 Collagen Gel Containing α-Elastin on Proliferation and Migration of Vascular Smooth Muscle and Endothelial Cells. Cardiovasc. Surg. 1997, 5 (2), 176–183. DOI:10.1016/S0967-2109(97)00004-5.

(68)

Kielty, C. M. Elastic Fibres in Health and Disease. Expert Rev. Mol. Med. 2006, 8 (19), 1–23. DOI:10.1017/S146239940600007X.

(69)

Sottile, J. Fibronectin Polymerization Regulates the Composition and Stability of Extracellular Matrix Fibrils and Cell-Matrix Adhesions. Mol. Biol. Cell 2002, 13 (10), 3546–3559. DOI:10.1091/mbc.E02-010048.

(70)

Martinez-Lemus, L. A.; Hill, M. A.; Meininger, G. A. The Plastic Nature of the Vascular Wall: A Continuum of Remodeling Events Contributing to Control of Arteriolar Diameter and Structure. Physiology 2009, 24 (1), 45–57. DOI:10.1152/physiol.00029.2008.

(71)

Liu, B.; Itoh, H.; Louie, O.; Kubota, K.; Kent, K. C. The Role of Phospholipase C and Phosphatidylinositol 3-Kinase in Vascular Smooth Muscle Cell Migration and Proliferation. J. Surg. Res. 2004, 120 (2), 256– 265. DOI:10.1016/j.jss.2003.12.015.

(72)

Majesky, M. W. Adventitia and Perivascular Cells. Arterioscler. Thromb. Vasc. Biol. 2015, 35 (8), e31–e35. DOI:10.1161/ATVBAHA.115.306088.


Page 58 of 85

Page 59 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(73)

Majesky, M. W.; Dong, X. R.; Hoglund, V.; Mahoney, W. M.; Daum, G. The Adventitia: A Dynamic Interface Containing Resident Progenitor Cells. Arterioscler. Thromb. Vasc. Biol. 2011, 31 (7), 1530–1539. DOI:10.1161/ATVBAHA.110.221549.

(74)

Majesky, M. W. Vascular Development. Arterioscler. Thromb. Vasc. Biol. 2018, 38 (3), e17–e24. DOI:10.1161/ATVBAHA.118.310223.

(75)

Psaltis, P. J.; Puranik, A. S.; Spoon, D. B.; Chue, C. D.; Hoffman, S. J.; Witt, T. A.; Delacroix, S.; Kleppe, L. S.; Mueske, C. S.; Pan, S.; et al. Characterization of a Resident Population of Adventitial Macrophage Progenitor Cells in Postnatal Vasculature. Circ. Res. 2014, 115 (3), 364–375. DOI:10.1161/CIRCRESAHA.115.303299.

(76)

Stenmark, K. R.; Yeager, M. E.; El Kasmi, K. C.; Nozik-Grayck, E.; Gerasimovskaya, E. V.; Li, M.; Riddle, S. R.; Frid, M. G. The Adventitia: Essential Regulator of Vascular Wall Structure and Function. Annu. Rev. Physiol. 2013, 75 (1), 23–47. DOI:10.1146/annurev-physiol-030212-183802.

(77)

Hu, Y.; Xu, Q. Adventitial Biology: Differentiation and Function. Arterioscler. Thromb. Vasc. Biol. 2011, 31 (7), 1523–1529. DOI:10.1161/ATVBAHA.110.221176.

(78)

Wörsdörfer, P.; Mekala, S. R.; Bauer, J.; Edenhofer, F.; Kuerten, S.; Ergün, S. The Vascular Adventitia: An Endogenous, Omnipresent Source of Stem Cells in the Body. Pharmacol. Ther. 2017, 171, 13–29. DOI:10.1016/j.pharmthera.2016.07.017.

(79)

McNeill, E.; Iqbal, A. J.; Jones, D.; Patel, J.; Coutinho, P.; Taylor, L.; Greaves, D. R.; Channon, K. M. Tracking Monocyte Recruitment and Macrophage Accumulation in Atherosclerotic Plaque Progression Using a Novel HCD68GFP/ApoE −/− Reporter Mouse—Brief ReportHighlights. Arterioscler. Thromb. Vasc. Biol. 2017, 37 (2), 258–263. DOI:10.1161/ATVBAHA.116.308367.

(80)

Wagenseil, J. E.; Mecham, R. P. Vascular Extracellular Matrix and Arterial Mechanics. Physiol. Rev. 2009, 89 (3), 957–989. DOI:10.1152/physrev.00041.2008.

(81)

Davies, P. F. Endothelial Mechanisms of Flow-Mediated Athero-Protection and Susceptibility. Circ. Res. 2007, 101 (1), 10–12. DOI:10.1161/CIRCRESAHA.107.156539.

(82)

Passerini, A. G.; Polacek, D. C.; Shi, C.; Francesco, N. M.; Manduchi, E.; Grant, G. R.; Pritchard, W. F.; Powell, S.; Chang, G. Y.; Stoeckert, C. J.; et al. Coexisting Proinflammatory and Antioxidative Endothelial Transcription Profiles in a Disturbed Flow Region of the Adult Porcine Aorta. Proc. Natl. Acad. Sci. 2004,



101 (8), 2482–2487. DOI:10.1073/pnas.0305938101. (83)

Taylor, C. A.; Hughes, T. J.; Zarins, C. K. Finite Element Modeling of Three-Dimensional Pulsatile Flow in the Abdominal Aorta: Relevance to Atherosclerosis. Ann. Biomed. Eng. 1998, 26 (6), 975–987. DOI:10.1114/1.140.

(84)

Glagov, S.; Zarins, C.; Giddens, D. P.; Ku, D. N. Hemodynamics and Atherosclerosis. Insights and Perspectives Gained from Studies of Human Arteries. Arch. Pathol. Lab. Med. 1988, 112 (10), 1018–1031.

(85)

Ku, D. N. Blood Flow in Arteries. Annu. Rev. Fluid Mech. 1997, 29 (1), 399–434. DOI:10.1146/annurev.fluid.29.1.399.

(86)

Himburg, H. A.; Grzybowski, D. M.; Hazel, A. L.; LaMack, J. A.; Li, X.-M.; Friedman, M. H. Spatial Comparison between Wall Shear Stress Measures and Porcine Arterial Endothelial Permeability. Am. J. Physiol. Circ. Physiol. 2004, 286 (5), H1916–H1922. DOI:10.1152/ajpheart.00897.2003.

(87)

Hoskins, P. R.; Hardman, D. Three-Dimensional Imaging and Computational Modelling for Estimation of Wall Stresses in Arteries. Br. J. Radiol. 2009, 82 Spec No (SPEC. ISSUE 1), S3-17. DOI:10.1259/bjr/96847348.

(88)

Samet, M. M.; Lelkes, P. I. The Hemodynamic Environment of Endothelium In Vitro and Its Simulation In Vitro. In Mechanical Forces and the Endothelium; Lelkes, P. I., Ed.; Taylor & Francis: The Netherlands, 1992; pp 1–32.

(89)

Ku, D. N.; Giddens, D. P.; Zarins, C. K.; Glagov, S. Pulsatile Flow and Atherosclerosis in the Human Carotid Bifurcation. Positive Correlation between Plaque Location and Low Oscillating Shear Stress. Arterioscler. Thromb. Vasc. Biol. 1985, 5 (3), 293–302. DOI:10.1161/01.ATV.5.3.293.

(90)

Peiffer, V.; Sherwin, S. J.; Weinberg, P. D. Computation in the Rabbit Aorta of a New Metric – the Transverse Wall Shear Stress – to Quantify the Multidirectional Character of Disturbed Blood Flow. J. Biomech. 2013, 46 (15), 2651–2658. DOI:10.1016/j.jbiomech.2013.08.003.

(91)

Chakraborty, A.; Chakraborty, S.; Jala, V. R.; Haribabu, B.; Sharp, M. K.; Berson, R. E. Effects of Biaxial Oscillatory Shear Stress on Endothelial Cell Proliferation and Morphology. Biotechnol. Bioeng. 2012, 109 (3), 695–707. DOI:10.1002/bit.24352.

(92)

Ojha, M. Wall Shear Stress Temporal Gradient and Anastomotic Intimal Hyperplasia. Circ. Res. 1994, 74 (6), 1227–1231. DOI:10.1161/01.RES.74.6.1227.


Page 60 of 85

Page 61 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(93)

Lei, M.; Kleinstreuer, C.; Truskey, G. A. Numerical Investigation and Prediction of Atherogenic Sites in Branching Arteries. J. Biomech. Eng. 1995, 117 (3), 350. DOI:10.1115/1.2794191.

(94)

P. W. Longest, C. K. Computational Haemodynamics Analysis and Comparison Study of Arterio-Venous Grafts. J. Med. Eng. Technol. 2000, 24 (3), 102–110. DOI:10.1080/03091900050135013.

(95)

Hyun, S.; Kleinstreuer, C.; Archie, J. P. Hemodynamics Analyses of Arterial Expansions with Implications to Thrombosis and Restenosis. Med. Eng. Phys. 2000, 22 (1), 13–27. DOI:10.1016/S1350-4533(00)00006-0.

(96)

Goubergrits, L.; Affeld, K.; Fernandez-Britto, J.; Falcon, L. Atherosclerosis and Flow in Carotid Arteries with Authentic Geometries. Biorheology 2002, 39 (3–4), 519–524.

(97)

Himburg, H. A.; Friedman, M. H. Correspondence of Low Mean Shear and High Harmonic Content in the Porcine Iliac Arteries. J. Biomech. Eng. 2006, 128 (6), 852. DOI:10.1115/1.2354211.

(98)

Mantha, A.; Karmonik, C.; Benndorf, G.; Strother, C.; Metcalfe, R. Hemodynamics in a Cerebral Artery before and after the Formation of an Aneurysm. AJNR. Am. J. Neuroradiol. 2006, 27 (5), 1113–1118.

(99)

Huo, Y.; Guo, X.; Kassab, G. S. The Flow Field along the Entire Length of Mouse Aorta and Primary Branches. Ann. Biomed. Eng. 2008, 36 (5), 685–699. DOI:10.1007/s10439-008-9473-4.

(100)

Kaunas, R.; Nguyen, P.; Usami, S.; Chien, S. Cooperative Effects of Rho and Mechanical Stretch on Stress Fiber Organization. Proc. Natl. Acad. Sci. U. S. A. 2005, 102 (44), 15895–15900. DOI:10.1073/pnas.0506041102.

(101)

Mechanical Forces and the Endothelium, 1st ed.; Lelkes, P., Gimbrone, M. A., Eds.; CRC Press: London, 2003.

(102)

Isnard, R. N.; Pannier, B. M.; Laurent, S.; London, G. M.; Diebold, B.; Safar, M. E. Pulsatile Diameter and Elastic Modulus of the Aortic Arch in Essential Hypertension: A Noninvasive Study. J. Am. Coll. Cardiol. 1989, 13 (2), 399–405. DOI:10.1016/0735-1097(89)90518-4.

(103)

Anwar, M. A.; Shalhoub, J.; Lim, C. S.; Gohel, M. S.; Davies, A. H. The Effect of Pressure-Induced Mechanical Stretch on Vascular Wall Differential Gene Expression. J. Vasc. Res. 2012, 49 (6), 463–478. DOI:10.1159/000339151.

(104)

Benetos, A.; Laurent, S.; Hoeks, A. P.; Boutouyrie, P. H.; Safar, M. E. Arterial Alterations with Aging and High Blood Pressure. Arterioscler.Thromb. 1993, 13, 90–97.

(105)

Benetos, A.; Laurent, S.; Boutouyrie, P.; Safar, M. Alteration in the Carotid Artery Wall Properties with



Ageing and High Blood Pressure Level. J. Hypertens. Suppl. 1991, 9 (6), S112-3. (106)

Boutouyrie, P.; Laurent, S.; Benetos, A.; Girerd, X. J.; Hoeks, A. P.; Safar, M. E. Opposing Effects of Ageing on Distal and Proximal Large Arteries in Hypertensives. J. Hypertens. Suppl. 1992, 10 (6), S87-91.

(107)

O’Rourke, M. Mechanical Principles in Arterial Disease. Hypertension 1995, 26 (1), 2–9. DOI:10.1161/01.HYP.26.1.2.

(108)

Dobrin, P. B. Mechanical Properties of Arteries. Physiol. Rev. 1978, 58 (2), 397–460. DOI:10.1152/physrev.1978.58.2.397.

(109)

Thubrikar, M. J.; Roskelley, S. K.; Eppink, R. T. Study of Stress Concentration in the Walls of the Bovine Coronary Arterial Branch. J. Biomech. 1990, 23 (1), 15–26. DOI:10.1016/0021-9290(90)90365-A.

(110)

Thubrikar, M. J.; Robicsek, F. Pressure-Induced Arterial Wall Stress and Atherosclerosis. Ann. Thorac. Surg. 1995, 59 (6), 1594–1603. DOI:10.1016/0003-4975(94)01037-D.

(111)

Liu, X. M.; Ensenat, D.; Wang, H.; Schafer, A. I.; Durante, W. Physiologic Cyclic Stretch Inhibits Apoptosis in Vascular Endothelium. FEBS Lett. 2003, 541 (1–3), 52–56. DOI:10.1016/S00145793(03)00285-0.

(112)

Zhao, S. Z.; Xu, X. Y.; Hughes, A. D.; Thom, S. A.; Stanton, A. V.; Ariff, B.; Long, Q. Blood Flow and Vessel Mechanics in a Physiologically Realistic Model of a Human Carotid Arterial Bifurcation. J. Biomech. 2000, 33 (8), 975–984. DOI:10.1016/S0021-9290(00)00043-9.

(113)

Zhao, S. Z.; Xu, X. Y.; Collins, M. W. The Numerical Analysis of Fluid-Solid Interactions for Blood Flow in Arterial Structures Part 2: Development of Coupled Fluid-Solid Algorithms. Proc. Inst. Mech. Eng. Part H J. Eng. Med. 1998, 212 (4), 241–252. DOI:10.1243/0954411981534024.

(114)

Kim, Y.-H.; Kim, J.-E.; Ito, Y.; Shih, A. M.; Brott, B.; Anayiotos, A. Hemodynamic Analysis of a Compliant Femoral Artery Bifurcation Model Using a Fluid Structure Interaction Framework. Ann. Biomed. Eng. 2008, 36 (11), 1753–1763. DOI:10.1007/s10439-008-9558-0.

(115)

Fraser, K. H.; Li, M.-X.; Lee, W. T.; Easson, W. J.; Hoskins, P. R. Fluid—structure Interaction in Axially Symmetric Models of Abdominal Aortic Aneurysms. Proc. Inst. Mech. Eng. Part H J. Eng. Med. 2009, 223 (2), 195–209. DOI:10.1243/09544119JEIM443.

(116)

Leung, J. H.; Wright, A. R.; Cheshire, N.; Crane, J.; Thom, S. A.; Hughes, A. D.; Xu, Y. Fluid Structure Interaction of Patient Specific Abdominal Aortic Aneurisms: A Comparison with Solid Stress Models.


Page 62 of 85

Page 63 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Biomed. Eng. Online 2006, 5, 1–15. DOI:10.1186/1475-925X-5-33. (117)

Tada, S.; Tarbell, J. M. Hemodynamics in Physio- and Pathological Vessels. In Vascular Engineering; Tanishita, K., Yamamoto, K., Eds.; Springer Japan: Tokyo, 2016; pp 69–98. DOI:10.1007/978-4-43154801-0.

(118)

Guyton, A. C.; Hall, J. E. Vascular Distensibility and Functions of the Arterial and Venous Systems. In Textbook of Medical Physiology; Elsevier: Philadelphia, PA, 2006; pp 171–180.

(119)

Guyton, A. C. The Relationship of Cardiac Output and Arterial Pressure Control. Circulation 1981, 64 (6), 1079–1088. DOI:10.1161/01.CIR.64.6.1079.

(120)

Guyton, A. C.; Hall, J. E. Overview of the Circulation; Medical Physics of Pressure, Flow, and Resistance. In Textbook of Medical Physiology; Elsevier: Philadelphia, PA, 2006; pp 161–180.

(121)

Whelton, P. K.; Carey, R. M.; Aronow, W. S.; Casey, D. E.; Collins, K. J.; Dennison Himmelfarb, C.; DePalma, S. M.; Gidding, S.; Jamerson, K. A.; Jones, D. W.; et al. 2017 ACC/AHA/AAPA/ABC/ACPM/AGS/APhA/ASH/ASPC/NMA/PCNA Guideline for the Prevention, Detection, Evaluation, and Management of High Blood Pressure in Adults. J. Am. Coll. Cardiol. 2018, 71 (19), e127–e248. DOI:10.1016/j.jacc.2017.11.006.

(122)

Wood, J. A.; Liliensiek, S. J.; Russell, P.; Nealey, P. F.; Murphy, C. J. Biophysical Cueing and Vascular Endothelial Cell Behavior. Materials (Basel). 2010, 3 (3), 1620–1639. DOI:10.3390/ma3031620.

(123)

Jacot, J. G.; Dianis, S.; Schnall, J.; Wong, J. Y. A Simple Microindentation Technique for Mapping the Microscale Compliance of Soft Hydrated Materials and Tissues. J. Biomed. Mater. Res. Part A 2006, 79A (3), 485–494. DOI:10.1002/jbm.a.30812.

(124)

Oie, T.; Murayama, Y.; Fukuda, T.; Nagai, C.; Omata, S.; Kanda, K.; Yaku, H.; Nakayama, Y. Local Elasticity Imaging of Vascular Tissues Using a Tactile Mapping System. J. Artif. Organs 2009, 12 (1), 40– 46. DOI:10.1007/s10047-008-0440-5.

(125)

Kesava Reddy, G. AGE-Related Cross-Linking of Collagen Is Associated with Aortic Wall Matrix Stiffness in the Pathogenesis of Drug-Induced Diabetes in Rats. Microvasc. Res. 2004, 68 (2), 132–142. DOI:10.1016/j.mvr.2004.04.002.

(126)

Assoul, N.; Flaud, P.; Chaouat, M.; Letourneur, D.; Bataille, I. Mechanical Properties of Rat Thoracic and Abdominal Aortas. J. Biomech. 2008, 41 (10), 2227–2236. DOI:10.1016/j.jbiomech.2008.04.017.



(127)

McKee, C. T.; Last, J. A.; Russell, P.; Murphy, C. J. Indentation Versus Tensile Measurements of Young’s Modulus for Soft Biological Tissues. Tissue Eng. Part B Rev. 2011, 17 (3), 155–164. DOI:10.1089/ten.teb.2010.0520.

(128)

Akhtar, R.; Sherratt, M. J.; Cruickshank, J. K.; Derby, B. Characterizing the Elastic Properties of Tissues. Mater. Today 2011, 14 (3), 96–105. DOI:10.1016/S1369-7021(11)70059-1.

(129)

Sasaki, N.; Odajima, S. Elongation Mechanism of Collagen Fibrils and Force-Strain Relations of Tendon at Each Level of Structural Hierarchy. J. Biomech. 1996, 29 (9), 1131–1136. DOI:10.1016/00219290(96)00024-3.

(130)

Harley, R.; James, D.; Miller, A.; White, J. W. Phonons and the Elastic Moduli of Collagen and Muscle. Nature 1977, 267 (5608), 285–287. DOI:10.1038/267285a0.

(131)

Gautieri, A.; Vesentini, S.; Redaelli, A.; Buehler, M. J. Viscoelastic Properties of Model Segments of Collagen Molecules. Matrix Biol. 2012, 31 (2), 141–149. DOI:10.1016/j.matbio.2011.11.005.

(132)

Shen, Z. L.; Dodge, M. R.; Kahn, H.; Ballarini, R.; Eppell, S. J. Stress-Strain Experiments on Individual Collagen Fibrils. Biophys. J. 2008, 95 (8), 3956–3963. DOI:10.1529/biophysj.107.124602.

(133)

Heim, A. J.; Matthews, W. G.; Koob, T. J. Determination of the Elastic Modulus of Native Collagen Fibrils via Radial Indentation. Appl. Phys. Lett. 2006, 89 (18). DOI:10.1063/1.2367660.

(134)

Lundkvist, A.; Lilleodden, E.; Siekhaus, W.; Kinney, J.; Pruitt, L.; Balooch, M. Viscoelastic Properties of Healthy Human Artery Measured in Saline Solution by AFM-Based Indentation Technique. MRS Proc. 1996, 436, 353. DOI:10.1557/PROC-436-353.

(135)

Wang, Y.; Hahn, J.; Zhang, Y. Mechanical Properties of Arterial Elastin With Water Loss. J. Biomech. Eng. 2018, 140 (4), 041012. DOI:10.1115/1.4038887.

(136)

Guarnieri, D.; Battista, S.; Borzacchiello, A.; Mayol, L.; De Rosa, E.; Keene, D. R.; Muscariello, L.; Barbarisi, A.; Netti, P. A. Effects of Fibronectin and Laminin on Structural, Mechanical and Transport Properties of 3D Collageneous Network. J. Mater. Sci. Mater. Med. 2007, 18 (2), 245–253. DOI:10.1007/s10856-006-0686-5.

(137)

Wen, Q.; Janmey, P. A. Effects of Non-Linearity on Cell-ECM Interactions. Exp. Cell Res. 2013, 319 (16), 2481–2489. DOI:10.1016/j.yexcr.2013.05.017.

(138)

Welling, L.; Zupka, M.; Welling, D. Mechanical Properties of Basement Membrane. Physiology 1995, 10


Page 64 of 85

Page 65 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(1), 30–35. DOI:10.1152/physiologyonline.1995.10.1.30. (139)

Gasiorowski, J. Z.; Murphy, C. J.; Nealey, P. F. Biophysical Cues and Cell Behavior: The Big Impact of Little Things. Annu. Rev. Biomed. Eng. 2013, 15 (1), 155–176. DOI:10.1146/annurev-bioeng-071811150021.

(140)

LeBleu, V. S.; Macdonald, B.; Kalluri, R. Structure and Function of Basement Membranes. Exp. Biol. Med. (Maywood). 2007, 232 (9), 1121–1129. DOI:10.3181/0703-MR-72.

(141)

Liliensiek, S. J.; Nealey, P.; Murphy, C. J. Characterization of Endothelial Basement Membrane Nanotopography in Rhesus Macaque as a Guide for Vessel Tissue Engineering. Tissue Eng Part A 2009, 15 (9), 2643–2651. DOI:10.1089/ten.TEA.2008.0284.

(142)

Harloff, A.; Nußbaumer, A.; Bauer, S.; Stalder, A. F.; Frydrychowicz, A.; Weiller, C.; Hennig, J.; Markl, M. In Vivo Assessment of Wall Shear Stress in the Atherosclerotic Aorta Using Flow-Sensitive 4D MRI. Magn. Reson. Med. 2010, 63 (6), 1529–1536. DOI:10.1002/mrm.22383.

(143)

von Knobelsdorff-Brenkenhoff, F.; Karunaharamoorthy, A.; Trauzeddel, R. F.; Barker, A. J.; Blaszczyk, E.; Markl, M.; Schulz-Menger, J. Evaluation of Aortic Blood Flow and Wall Shear Stress in Aortic Stenosis and Its Association With Left Ventricular Remodeling. Circ. Cardiovasc. Imaging 2016, 9 (3), e004038. DOI:10.1161/CIRCIMAGING.115.004038.

(144)

Yang, J.; Cho, K.; Kim, J.; Kim, S.; Kim, C.; You, G.; Lee, J.; Choi, S.; Lee, S.; Kim, H.; et al. Wall Shear Stress in Hypertensive Patients Is Associated with Carotid Vascular Deformation Assessed by Speckle Tracking Strain Imaging. Clin. Hypertens. 2014, 20 (1), 10. DOI:10.1186/2056-5909-20-10.

(145)

Schäfer, M.; Kheyfets, V. O.; Schroeder, J. D.; Dunning, J.; Shandas, R.; Buckner, J. K.; Browning, J.; Hertzberg, J.; Hunter, K. S.; Fenster, B. E. Main Pulmonary Arterial Wall Shear Stress Correlates with Invasive Hemodynamics and Stiffness in Pulmonary Hypertension. Pulm. Circ. 2016, 6 (1), 37–45. DOI:10.1086/685024.

(146)

Xi, C.; Latnie, C.; Zhao, X.; Tan, J. Le; Wall, S. T.; Genet, M.; Zhong, L.; Lee, L. C. Patient-Specific Computational Analysis of Ventricular Mechanics in Pulmonary Arterial Hypertension. J. Biomech. Eng. 2016, 138 (11), 111001. DOI:10.1115/1.4034559.

(147)

Lai, Y.-C.; Potoka, K. C.; Champion, H. C.; Mora, A. L.; Gladwin, M. T. Pulmonary Arterial Hypertension: The Clinical Syndrome. Circ. Res. 2014, 115 (1), 115–130. DOI:10.1161/CIRCRESAHA.115.301146.



(148)

Galizia, M. S.; Barker, A.; Liao, Y.; Collins, J.; Carr, J.; McDermott, M. M.; Markl, M. Wall Morphology, Blood Flow and Wall Shear Stress: MR Findings in Patients with Peripheral Artery Disease. Eur. Radiol. 2014, 24 (4), 850–856. DOI:10.1007/s00330-013-3081-x.

(149)

Benetos, A.; Laurent, S.; Hoeks, A. P.; Boutouyrie, P. H.; Safar, M. E. Arterial Alterations with Aging and High Blood Pressure. A Noninvasive Study of Carotid and Femoral Arteries. Arterioscler. Thromb. a J. Vasc. Biol. 1993, 13 (1), 90–97. DOI:10.1161/atv91.13.1.8422344.

(150)

Suurküla, M.; Fagerberg, B.; Wendelhag, I.; Agewall, S.; Wikstrand, J. Atherosclerotic Disease in the Femoral Artery in Hypertensive Patients at High Cardiovascular Risk. The Value of Ultrasonographic Assessment of Intima-Media Thickness and Plaque Occurrence. Risk Intervention Study (RIS) Group. Arterioscler. Thromb. Vasc. Biol. 1996, 16 (8), 971–977. DOI:10.1161/atvb.16.8.971.

(151)

Hsiai, T. K.; Blackman, B.; Jo, H. Hemodynamics and Mechanobiology of Endothelium; Hsiai, T. K., Blackman, B., Jo, H., Eds.; WORLD SCIENTIFIC: Singapore, 2010. DOI:10.1142/7385.

(152)

Cellular Mechanotransduction; Mofrad, M. R. K., Kamm, R. D., Eds.; Cambridge University Press: Cambridge, 2009. DOI:10.1017/CBO9781139195874.

(153)

Davies, P. F. Flow-Mediated Endothelial Mechanotransduction. Physiol. Rev. 1995, 75 (3), 519–560. DOI:10.1152/physrev.1995.75.3.519.

(154)

Cooper, G. M.; Hausman, R. E. The Cell: A Molecular Approach, 7th ed.; Sinauer, 2016.

(155)

Butler, P. J.; Chien, S. Role of the Plasma Membrane in Endothelial Cell Mechanosensation of Shear Stress. In Cellular Mechanotransduction; Mofrad, M. R. K., Kamm, R. D., Eds.; Cambridge University Press: Cambridge, 2009; pp 61–88. DOI:10.1017/CBO9781139195874.

(156)

Yamamoto, K.; Ando, J. Vascular Endothelial Cell Membranes Differentiate between Stretch and Shear Stress through Transitions in Their Lipid Phases. Am. J. Physiol. Circ. Physiol. 2015, 309 (7), H1178– H1185. DOI:10.1152/ajpheart.00241.2015.

(157)

Maniotis, A. J.; Chen, C. S.; Ingber, D. E. Demonstration of Mechanical Connections between Integrins, Cytoskeletal Filaments, and Nucleoplasm That Stabilize Nuclear Structure. Proc. Natl. Acad. Sci. 1997, 94 (3), 849–854. DOI:10.1073/pnas.94.3.849.

(158)

Guilluy, C.; Burridge, K. Nuclear Mechanotransduction: Forcing the Nucleus to Respond. Nucleus 2015, 6 (1), 19–22. DOI:10.1080/19491034.2014.1001705.


Page 66 of 85

Page 67 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(159)

Dahl, K. N.; Ribeiro, A. J. S.; Lammerding, J. Nuclear Shape, Mechanics, and Mechanotransduction. Circ. Res. 2008, 102 (11), 1307–1318. DOI:10.1161/CIRCRESAHA.108.173989.

(160)

Isermann, P.; Lammerding, J. Nuclear Mechanics and Mechanotransduction in Health and Disease. Curr. Biol. 2013, 23 (24), R1113–R1121. DOI:10.1016/j.cub.2013.11.009.

(161)

Cho, S.; Irianto, J.; Discher, D. E. Mechanosensing by the Nucleus: From Pathways to Scaling Relationships. J. Cell Biol. 2017, 216 (2), 305–315. DOI:10.1083/jcb.201610042.

(162)

Weinbaum, S.; Zhang, X.; Han, Y.; Vink, H.; Cowin, S. C. Mechanotransduction and Flow across the Endothelial Glycocalyx. Proc. Natl. Acad. Sci. 2003, 100 (13), 7988–7995. DOI:10.1073/pnas.1332808100.

(163)

Oike, M.; Watanabe, M.; Kimura, C. Involvement of Heparan Sulfate Proteoglycan in Sensing Hypotonic Stress in Bovine Aortic Endothelial Cells. Biochim. Biophys. Acta - Gen. Subj. 2008, 1780 (10), 1148–1155. DOI:10.1016/j.bbagen.2008.07.004.

(164)

Tarbell, J. M.; Cancel, L. M. The Glycocalyx and Its Significance in Human Medicine. J. Intern. Med. 2016, 280 (1), 97–113. DOI:10.1111/joim.12465.

(165)

Simon, M.; Strathmann, M.; Gautam, N. Diversity of G Proteins in Signal Transduction. Science (80-. ). 1991, 252 (5007), 802–808. DOI:10.1126/science.1902986.

(166)

Oldham, W. M.; Hamm, H. E. Heterotrimeric G Protein Activation by G-Protein-Coupled Receptors. Nat. Rev. Mol. Cell Biol. 2008, 9 (1), 60–71. DOI:10.1038/nrm2299.

(167)

Hsieh, H.-J.; Li, N.-Q.; Frangos, J. A. Pulsatile and Steady Flow Induces C-Fos Expression in Human Endothelial Cells. J. Cell. Physiol. 1993, 154 (1), 143–151. DOI:10.1002/jcp.1041540118.

(168)

Gudi, S. R.; Clark, C. B.; Frangos, J. A. Fluid Flow Rapidly Activates G Proteins in Human Endothelial Cells. Involvement of G Proteins in Mechanochemical Signal Transduction. Circ. Res. 1996, 79 (4), 834– 839. DOI:10.1161/res.79.4.834.

(169)

Gudi, S.; Nolan, J. P.; Frangos, J. A. Modulation of GTPase Activity of G Proteins by Fluid Shear Stress and Phospholipid Composition. Proc. Natl. Acad. Sci. 1998, 95 (5), 2515–2519. DOI:10.1073/pnas.95.5.2515.

(170)

Hein, P.; Frank, M.; Hoffmann, C.; Lohse, M. J.; Bünemann, M. Dynamics of Receptor/G Protein Coupling in Living Cells. EMBO J. 2005, 24 (23), 4106–4114. DOI:10.1038/sj.emboj.7600870.

(171)

Azpiazu, I.; Gautam, N. A Fluorescence Resonance Energy Transfer-Based Sensor Indicates That Receptor Access to a G Protein Is Unrestricted in a Living Mammalian Cell. J. Biol. Chem. 2004, 279 (26), 27709–



27718. DOI:10.1074/jbc.M403712200. (172)

Yan-Liang Zhang, J.; Frangos, ohn A.; Chachisvilis, M. Mechanotransduction by Membrane-Mediated Activation of G-Protein Coupled Receptors and G-Proteins. In Cellular Mechanotransduction; Mofrad, M. R. K., Kamm, R. D., Eds.; Cambridge University Press: Cambridge, 2009; pp 89–119. DOI:10.1017/CBO9781139195874.

(173)

Chachisvilis, M.; Zhang, Y.-L.; Frangos, J. A. G Protein-Coupled Receptors Sense Fluid Shear Stress in Endothelial Cells. Proc. Natl. Acad. Sci. 2006, 103 (42), 15463–15468. DOI:10.1073/pnas.0607224103.

(174)

dela Paz, N. G.; Melchior, B.; Shayo, F. Y.; Frangos, J. A. Heparan Sulfates Mediate the Interaction between Platelet Endothelial Cell Adhesion Molecule-1 (PECAM-1) and the Gα q/11 Subunits of Heterotrimeric G Proteins. J. Biol. Chem. 2014, 289 (11), 7413–7424. DOI:10.1074/jbc.M113.542514.

(175)

Otte, L. A.; Bell, K. S.; Loufrani, L.; Yeh, J.-C.; Melchior, B.; Dao, D. N.; Stevens, H. Y.; White, C. R.; Frangos, J. A. Rapid Changes in Shear Stress Induce Dissociation of a Gα q/11 -Platelet Endothelial Cell Adhesion Molecule-1 Complex. J. Physiol. 2009, 587 (10), 2365–2373. DOI:10.1113/jphysiol.2009.172643.

(176)

Ranade, S. S.; Syeda, R.; Patapoutian, A. Mechanically Activated Ion Channels. Neuron 2015, 87 (6), 1162– 1179. DOI:10.1016/j.neuron.2015.08.032.

(177)

Murthy, S. E.; Dubin, A. E.; Patapoutian, A. Piezos Thrive under Pressure: Mechanically Activated Ion Channels in Health and Disease. Nat. Rev. Mol. Cell Biol. 2017, 18 (12), 771–783. DOI:10.1038/nrm.2017.92.

(178)

Barakat, A. I.; Gojova, A. Role of Ion Channels in Cellular Mechanotransduction – Lessons from the Vascular Endothelium. In Cellular Mechanotransduction; Mofrad, M. R. K., Kamm, R. D., Eds.; Cambridge University Press: Cambridge, 2009; pp 161–180. DOI:10.1017/CBO9781139195874.

(179)

Lieu, D. K.; Pappone, P. A.; Barakat, A. I. Differential Membrane Potential and Ion Current Responses to Different Types of Shear Stress in Vascular Endothelial Cells. Am. J. Physiol. Physiol. 2004, 286 (6), C1367–C1375. DOI:10.1152/ajpcell.00243.2003.

(180)

Gautam, M.; Shen, Y.; Thirkill, T. L.; Douglas, G. C.; Barakat, A. I. Flow-Activated Chloride Channels in Vascular Endothelium. J. Biol. Chem. 2006, 281 (48), 36492–36500. DOI:10.1074/jbc.M605866200.

(181)

Barakat, A. I.; Lieu, D. K.; Gojova, A. Secrets of the Code: Do Vascular Endothelial Cells Use Ion Channels to Decipher Complex Flow Signals? Biomaterials 2006, 27 (5), 671–678.


Page 68 of 85

Page 69 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


DOI:10.1016/j.biomaterials.2005.07.036. (182)

Ranade, S. S.; Qiu, Z.; Woo, S.-H.; Hur, S. S.; Murthy, S. E.; Cahalan, S. M.; Xu, J.; Mathur, J.; Bandell, M.; Coste, B.; et al. Piezo1, a Mechanically Activated Ion Channel, Is Required for Vascular Development in Mice. Proc. Natl. Acad. Sci. 2014, 111 (28), 10347–10352. DOI:10.1073/pnas.1409233111.

(183)

Coste, B.; Mathur, J.; Schmidt, M.; Earley, T. J.; Ranade, S.; Petrus, M. J.; Dubin, A. E.; Patapoutian, A. Piezo1 and Piezo2 Are Essential Components of Distinct Mechanically Activated Cation Channels. Science (80-. ). 2010, 330 (6000), 55–60. DOI:10.1126/science.1193270.

(184)

Volkers, L.; Mechioukhi, Y.; Coste, B. Piezo Channels: From Structure to Function. Pflügers Arch. - Eur. J. Physiol. 2015, 467 (1), 95–99. DOI:10.1007/s00424-014-1578-z.

(185)

Rode, B.; Shi, J.; Endesh, N.; Drinkhill, M. J.; Webster, P. J.; Lotteau, S. J.; Bailey, M. A.; Yuldasheva, N. Y.; Ludlow, M. J.; Cubbon, R. M.; et al. Piezo1 Channels Sense Whole Body Physical Activity to Reset Cardiovascular Homeostasis and Enhance Performance. Nat. Commun. 2017, 8 (1), 350. DOI:10.1038/s41467-017-00429-3.

(186)

Beech, D. J. Endothelial Piezo1 Channels as Sensors of Exercise. J. Physiol. 2018, 596 (6), 979–984. DOI:10.1113/JP274396.

(187)

Sackin, H. Stretch-Activated Ion Channels. Kidney Int. 1995, 48 (4), 1134–1147. DOI:10.1038/ki.1995.397.

(188)

Thodeti, C. K.; Matthews, B.; Ravi, A.; Mammoto, A.; Ghosh, K.; Bracha, A. L.; Ingber, D. E. TRPV4 Channels Mediate Cyclic Strain-Induced Endothelial Cell Reorientation Through Integrin-to-Integrin Signaling. Circ. Res. 2009, 104 (9), 1123–1130. DOI:10.1161/CIRCRESAHA.108.192930.

(189)

Naruse, K.; Yamada, T.; Sokabe, M. Involvement of SA Channels in Orienting Response of Cultured Endothelial Cells to Cyclic Stretch. Am. J. Physiol. Circ. Physiol. 1998, 274 (5), H1532–H1538. DOI:10.1152/ajpheart.1998.274.5.H1532.

(190)

Lansman, J. B.; Hallam, T. J.; Rink, T. J. Single Stretch-Activated Ion Channels in Vascular Endothelial Cells as Mechanotransducers? Nature 1987, 325 (6107), 811–813. DOI:10.1038/325811a0.

(191)

Doyle, A. D.; Yamada, K. M. Cellular Mechanotransduction: Interactions with the Extracellular Matrix. In Cellular Mechanotransduction; Mofrad, M. R. K., Kamm, R. D., Eds.; Cambridge University Press: Cambridge, 2009; pp 120–160. DOI:10.1017/CBO9781139195874.

(192)

Ruoslahti, E. RGD and Other Recognition Sequences for Integrins. Annu. Rev. Cell Dev. Biol. 1996, 12 (1),



697–715. DOI:10.1146/annurev.cellbio.12.1.697. (193)

Winograd-Katz, S. E.; Fässler, R.; Geiger, B.; Legate, K. R. The Integrin Adhesome: From Genes and Proteins to Human Disease. Nat. Rev. Mol. Cell Biol. 2014, 15 (4), 273–288. DOI:10.1038/nrm3769.

(194)

Geiger, B.; Spatz, J. P.; Bershadsky, A. D. Environmental Sensing through Focal Adhesions. Nat. Rev. Mol. Cell Biol. 2009, 10 (1), 21–33. DOI:10.1038/nrm2593.

(195)

Carter, S. B. Haptotaxis and the Mechanism of Cell Motility. Nature 1967, 213 (5073), 256–260. DOI:10.1038/213256a0.

(196)

Cavalcanti-Adam, E. A.; Volberg, T.; Micoulet, A.; Kessler, H.; Geiger, B.; Spatz, J. P. Cell Spreading and Focal Adhesion Dynamics Are Regulated by Spacing of Integrin Ligands. Biophys. J. 2007, 92 (8), 2964– 2974. DOI:10.1529/biophysj.106.089730.

(197)

Duband, J. L. Fibronectin Receptor Exhibits High Lateral Mobility in Embryonic Locomoting Cells but Is Immobile in Focal Contacts and Fibrillar Streaks in Stationary Cells. J. Cell Biol. 1988, 107 (4), 1385–1396. DOI:10.1083/jcb.107.4.1385.

(198)

Lo, C.-M.; Wang, H.-B.; Dembo, M.; Wang, Y. Cell Movement Is Guided by the Rigidity of the Substrate. Biophys. J. 2000, 79 (1), 144–152. DOI:10.1016/S0006-3495(00)76279-5.

(199)

Gong, Z.; Szczesny, S. E.; Caliari, S. R.; Charrier, E. E.; Chaudhuri, O.; Cao, X.; Lin, Y.; Mauck, R. L.; Janmey, P. A.; Burdick, J. A.; et al. Matching Material and Cellular Timescales Maximizes Cell Spreading on Viscoelastic Substrates. Proc. Natl. Acad. Sci. 2018, 115 (12), E2686–E2695. DOI:10.1073/pnas.1716620115.

(200)

Tusan, C. G.; Man, Y.-H.; Zarkoob, H.; Johnston, D. A.; Andriotis, O. G.; Thurner, P. J.; Yang, S.; Sander, E. A.; Gentleman, E.; Sengers, B. G.; et al. Collective Cell Behavior in Mechanosensing of Substrate Thickness. Biophys. J. 2018, 114 (11), 2743–2755. DOI:10.1016/j.bpj.2018.03.037.

(201)

Weiss, P. Experiments on Cell and Axon Orientation in Vitro: The Role of Colloidal Exudates in Tissue Organization. J. Exp. Zool. 1945, 100 (3), 353–386. DOI:10.1002/jez.1401000305.

(202)

Unadkat, H. V.; Hulsman, M.; Cornelissen, K.; Papenburg, B. J.; Truckenmuller, R. K.; Carpenter, A. E.; Wessling, M.; Post, G. F.; Uetz, M.; Reinders, M. J. T.; et al. An Algorithm-Based Topographical Biomaterials Library to Instruct Cell Fate. Proc. Natl. Acad. Sci. 2011, 108 (40), 16565–16570. DOI:10.1073/pnas.1109861108.


Page 70 of 85

Page 71 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(203)

Bettinger, C. J.; Langer, R.; Borenstein, J. T. Engineering Substrate Topography at the Micro- and Nanoscale to Control Cell Function. Angew. Chemie Int. Ed. 2009, 48 (30), 5406–5415. DOI:10.1002/anie.200805179.

(204)

Kirschner, C. M.; Schumacher, J. F.; Brennan, A. B. Cellular Responses to Bio-Inspired Engineered Topography. In Bio-inspired Materials for Biomedical Engineering; John Wiley & Sons, Inc.: Hoboken, NJ, USA, 2014; pp 77–97. DOI:10.1002/9781118843499.ch5.

(205)

Dejana, E. Endothelial Cell–cell Junctions: Happy Together. Nat. Rev. Mol. Cell Biol. 2004, 5 (4), 261–270. DOI:10.1038/nrm1357.

(206)

Dorland, Y. L.; Huveneers, S. Cell–cell Junctional Mechanotransduction in Endothelial Remodeling. Cell. Mol. Life Sci. 2017, 74 (2), 279–292. DOI:10.1007/s00018-016-2325-8.

(207)

Huveneers, S.; Oldenburg, J.; Spanjaard, E.; van der Krogt, G.; Grigoriev, I.; Akhmanova, A.; Rehmann, H.; de Rooij, J. Vinculin Associates with Endothelial VE-Cadherin Junctions to Control Force-Dependent Remodeling. J. Cell Biol. 2012, 196 (5), 641–652. DOI:10.1083/jcb.201108120.

(208)

Tzima, E.; Irani-Tehrani, M.; Kiosses, W. B.; Dejana, E.; Schultz, D. A.; Engelhardt, B.; Cao, G.; DeLisser, H.; Schwartz, M. A. A Mechanosensory Complex That Mediates the Endothelial Cell Response to Fluid Shear Stress. Nature 2005, 437 (7057), 426–431. DOI:10.1038/nature03952.

(209)

Li, Y.-S. J.; Haga, J. H.; Chien, S. Molecular Basis of the Effects of Shear Stress on Vascular Endothelial Cells. J. Biomech. 2005, 38 (10), 1949–1971. DOI:10.1016/j.jbiomech.2004.09.030.

(210)

Collins, C.; Guilluy, C.; Welch, C.; O’Brien, E. T.; Hahn, K.; Superfine, R.; Burridge, K.; Tzima, E. Localized Tensional Forces on PECAM-1 Elicit a Global Mechanotransduction Response via the IntegrinRhoA Pathway. Curr. Biol. 2012, 22 (22), 2087–2094. DOI:10.1016/j.cub.2012.08.051.

(211)

Ueki, Y.; Sakamoto, N.; Ohashi, T.; Sato, M. Morphological Responses of Vascular Endothelial Cells Induced by Local Stretch Transmitted Through Intercellular Junctions. Exp. Mech. 2009, 49 (1), 125–134. DOI:10.1007/s11340-008-9143-3.

(212)

Davies, P. F.; Helmke, B. P. Endothelial Mechanotransduction. In Cellular Mechanotransduction; Mofrad, M. R. K., Kamm, R. D., Eds.; Cambridge University Press: Cambridge, 2009; pp 20–60. DOI:10.1017/CBO9781139195874.

(213)

Dabagh, M.; Jalali, P.; Butler, P. J.; Randles, A.; Tarbell, J. M. Mechanotransmission in Endothelial Cells



Subjected to Oscillatory and Multi-Directional Shear Flow. J. R. Soc. Interface 2017, 14 (130), 20170185. DOI:10.1098/rsif.2017.0185. (214)

Ingber, D. E. Tensegrity as a Mechanism for Integrating Molecular and Cellular Mechanotransduction Mechanisms. In Cellular Mechanotransduction; Mofrad, M. R. K., Kamm, R. D., Eds.; Cambridge University Press: Cambridge, 2009; pp 196–219. DOI:10.1017/CBO9781139195874.

(215)

Zhou, J.; Li, Y.-S.; Chien, S. Shear Stress-Initiated Signaling and Its Regulation of Endothelial Function. Arterioscler. Thromb. Vasc. Biol. 2014, 34 (10), 2191–2198. DOI:10.1161/ATVBAHA.114.303422.

(216)

Zhou, J.; Li, Y.-S.; Chien, S. Shear Stress-Initiated Signaling and Its Regulation of Endothelial Function. Arterioscler. Thromb. Vasc. Biol. 2014, 34 (10), 2191–2198. DOI:10.1161/ATVBAHA.114.303422.

(217)

Romer, L. H. Focal Adhesions: Paradigm for a Signaling Nexus. Circ. Res. 2006, 98 (5), 606–616. DOI:10.1161/01.RES.0000207408.31270.db.

(218)

Atkins, G. B.; Jain, M. K. Role of Kruppel-Like Transcription Factors in Endothelial Biology. Circ. Res. 2007, 100 (12), 1686–1695. DOI:10.1161/01.RES.0000267856.00713.0a.

(219)

Wang, K.-C.; Yeh, Y.-T.; Nguyen, P.; Limqueco, E.; Lopez, J.; Thorossian, S.; Guan, K.-L.; Li, Y.-S. J.; Chien, S. Flow-Dependent YAP/TAZ Activities Regulate Endothelial Phenotypes and Atherosclerosis. Proc. Natl. Acad. Sci. 2016, 113 (41), 11525–11530. DOI:10.1073/pnas.1613121113.

(220)

Shyy, J. Y.-J. Role of Integrins in Endothelial Mechanosensing of Shear Stress. Circ. Res. 2002, 91 (9), 769–775. DOI:10.1161/01.RES.0000038487.19924.18.

(221)

Humphrey, J. D.; Schwartz, M. A.; Tellides, G.; Milewicz, D. M. Role of Mechanotransduction in Vascular Biology: Focus on Thoracic Aortic Aneurysms and Dissections. Circ. Res. 2015, 116 (8), 1448–1461. DOI:10.1161/CIRCRESAHA.114.304936.

(222)

Davies, P. F. Spatial Microstimuli in Endothelial Mechanosignaling. Circ. Res. 2003, 92 (4), 359–370. DOI:10.1161/01.RES.0000060201.41923.88.

(223)

Hahn, C.; Schwartz, M. A. The Role of Cellular Adaptation to Mechanical Forces in Atherosclerosis. Arterioscler. Thromb. Vasc. Biol. 2008, 28 (12), 2101–2107. DOI:10.1161/ATVBAHA.108.165951.

(224)

Gratton, J.-P. Caveolae and Caveolins in the Cardiovascular System. Circ. Res. 2004, 94 (11), 1408–1417. DOI:10.1161/01.RES.0000129178.56294.17.

(225)

Davis, C. A.; Zambrano, S.; Anumolu, P.; Allen, A. C. B.; Sonoqui, L.; Moreno, M. R. Device-Based In


Page 72 of 85

Page 73 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Vitro Techniques for Mechanical Stimulation of Vascular Cells: A Review. J. Biomech. Eng. 2015, 137 (4), 040801. DOI:10.1115/1.4029016. (226)

Sinha, R.; Verdonschot, N.; Koopman, B.; Rouwkema, J. Tuning Cell and Tissue Development by Combining Multiple Mechanical Signals. Tissue Eng. Part B Rev. 2017, 23 (5), 494–504. DOI:10.1089/ten.teb.2016.0500.

(227)

Gray, K. M.; Stroka, K. M. Vascular Endothelial Cell Mechanosensing: New Insights Gained from Biomimetic Microfluidic Models. Semin. Cell Dev. Biol. 2017, 71, 106–117. DOI:10.1016/j.semcdb.2017.06.002.

(228)

Chiu, J.-J.; Chien, S. Effects of Disturbed Flow on Vascular Endothelium: Pathophysiological Basis and Clinical Perspectives. Physiol. Rev. 2011, 91 (1), 327–387. DOI:10.1152/physrev.00047.2009.

(229)

Spruell, C.; Baker, A. B. Analysis of a High-Throughput Cone-and-Plate Apparatus for the Application of Defined Spatiotemporal Flow to Cultured Cells. Biotechnol. Bioeng. 2013, 110 (6), 1782–1793. DOI:10.1002/bit.24823.

(230)

Galie, P. A.; van Oosten, A.; Chen, C. S.; Janmey, P. A. Application of Multiple Levels of Fluid Shear Stress to Endothelial Cells Plated on Polyacrylamide Gels. Lab Chip 2015, 15 (4), 1205–1212. DOI:10.1039/C4LC01236D.

(231)

Dardik, A.; Chen, L.; Frattini, J.; Asada, H.; Aziz, F.; Kudo, F. A.; Sumpio, B. E. Differential Effects of Orbital and Laminar Shear Stress on Endothelial Cells. J. Vasc. Surg. 2005, 41 (5), 869–880. DOI:10.1016/j.jvs.2005.01.020.

(232)

Xu, J.; Mathur, J.; Vessières, E.; Hammack, S.; Nonomura, K.; Favre, J.; Grimaud, L.; Petrus, M.; Francisco, A.; Li, J.; et al. GPR68 Senses Flow and Is Essential for Vascular Physiology. Cell 2018, 173 (3), 762–775.e16. DOI:10.1016/j.cell.2018.03.076.

(233)

Kamble, H.; Barton, M. J.; Jun, M.; Park, S.; Nguyen, N.-T. Cell Stretching Devices as Research Tools: Engineering and Biological Considerations. Lab Chip 2016, 16 (17), 3193–3203. DOI:10.1039/C6LC00607H.

(234)

Yoshino, D.; Sato, K.; Sato, M. Endothelial Cell Response Under Hydrostatic Pressure Condition Mimicking Pressure Therapy. Cell. Mol. Bioeng. 2015, 8 (2), 296–303. DOI:10.1007/s12195-015-0385-8.

(235)

Liu, M.-C.; Shih, H.-C.; Wu, J.-G.; Weng, T.-W.; Wu, C.-Y.; Lu, J.-C.; Tung, Y.-C. Electrofluidic Pressure



Sensor Embedded Microfluidic Device: A Study of Endothelial Cells under Hydrostatic Pressure and Shear Stress Combinations. Lab Chip 2013, 13 (9), 1743. DOI:10.1039/c3lc41414k. (236)

Huveneers, S.; Daemen, M. J. A. P.; Hordijk, P. L. Between Rho(k) and a Hard Place: The Relation Between Vessel Wall Stiffness, Endothelial Contractility, and Cardiovascular Disease. Circ. Res. 2015, 116 (5), 895–908. DOI:10.1161/CIRCRESAHA.116.305720.

(237)

Chang, H.; Liu, X.; Hu, M.; Zhang, H.; Li, B.; Ren, K.; Boudou, T.; Albiges-Rizo, C.; Picart, C.; Ji, J. Substrate Stiffness Combined with Hepatocyte Growth Factor Modulates Endothelial Cell Behavior. Biomacromolecules 2016, 17 (9), 2767–2776. DOI:10.1021/acs.biomac.6b00318.

(238)

Lampi, M. C.; Guvendiren, M.; Burdick, J. A.; Reinhart-King, C. A. Photopatterned Hydrogels to Investigate the Endothelial Cell Response to Matrix Stiffness Heterogeneity. ACS Biomater. Sci. Eng. 2017, 3 (11), 3007–3016. DOI:10.1021/acsbiomaterials.6b00633.

(239)

Byfield, F. J.; Reen, R. K.; Shentu, T.-P.; Levitan, I.; Gooch, K. J. Endothelial Actin and Cell Stiffness Is Modulated by Substrate Stiffness in 2D and 3D. J. Biomech. 2009, 42 (8), 1114–1119. DOI:10.1016/j.jbiomech.2009.02.012.

(240)

Jeon, H.; Tsui, J. H.; Jang, S. I.; Lee, J. H.; Park, S.; Mun, K.; Boo, Y. C.; Kim, D.-H. Combined Effects of Substrate Topography and Stiffness on Endothelial Cytokine and Chemokine Secretion. ACS Appl. Mater. Interfaces 2015, 7 (8), 4525–4532. DOI:10.1021/acsami.5b00554.

(241)

Feinberg, A. W.; Wilkerson, W. R.; Seegert, C. A.; Gibson, A. L.; Hoipkemeier‐Wilson, L.; Brennan, A. B. Systematic Variation of Microtopography, Surface Chemistry and Elastic Modulus and the State Dependent Effect on Endothelial Cell Alignment. J. Biomed. Mater. Res. Part A 2008, 86A (2), 522–534. DOI:10.1002/jbm.a.31626.

(242)

Ding, Y.; Floren, M.; Tan, W. High-Throughput Screening of Vascular Endothelium-Destructive or Protective Microenvironments: Cooperative Actions of Extracellular Matrix Composition, Stiffness, and Structure. Adv. Healthc. Mater. 2017, 6 (11), 1601426. DOI:10.1002/adhm.201601426.

(243)

Kannan, R. Y.; Salacinski, H. J.; Sales, K.; Butler, P.; Seifalian, A. M. The Roles of Tissue Engineering and Vascularisation in the Development of Micro-Vascular Networks: A Review. Biomaterials 2005, 26 (14), 1857–1875. DOI:10.1016/j.biomaterials.2004.07.006.

(244)

Derricks, K. E.; Trinkaus-Randall, V.; Nugent, M. A. Extracellular Matrix Stiffness Modulates VEGF


Page 74 of 85

Page 75 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Calcium Signaling in Endothelial Cells: Individual Cell and Population Analysis. Integr. Biol. 2015, 7 (9), 1011–1025. DOI:10.1039/C5IB00140D. (245)

Canver, A. C.; Ngo, O.; Urbano, R. L.; Clyne, A. M. Endothelial Directed Collective Migration Depends on Substrate Stiffness via Localized Myosin Contractility and Cell-Matrix Interactions. J. Biomech. 2016, 49 (8), 1369–1380. DOI:10.1016/j.jbiomech.2015.12.037.

(246)

Moore, E. M.; West, J. L. Bioactive Poly(Ethylene Glycol) Acrylate Hydrogels for Regenerative Engineering. Regen. Eng. Transl. Med. 2018. DOI:10.1007/s40883-018-0074-y.

(247)

Greiner, A. M.; Sales, A.; Chen, H.; Biela, S. A.; Kaufmann, D.; Kemkemer, R. Nano- and Microstructured Materials for in Vitro Studies of the Physiology of Vascular Cells. Beilstein J. Nanotechnol. 2016, 7, 1620– 1641. DOI:10.3762/bjnano.7.155.

(248)

Davies, P. F. Hemodynamic Shear Stress and the Endothelium in Cardiovascular Pathophysiology. Nat. Clin. Pract. Cardiovasc. Med. 2009, 6 (1), 16–26. DOI:10.1038/ncpcardio1397.

(249)

Sato, M.; Nagayama, K.; Kataoka, N.; Sasaki, M.; Hane, K. Local Mechanical Properties Measured by Atomic Force Microscopy for Cultured Bovine Endothelial Cells Exposed to Shear Stress. J. Biomech. 2000, 33 (1), 127–135. DOI:10.1016/S0021-9290(99)00178-5.

(250)

Sato, M.; Levesque, M. J.; Nerem, R. M. Micropipette Aspiration of Cultured Bovine Aortic Endothelial Cells Exposed to Shear Stress. Arteriosclerosis 1987, 7 (3), 276–286. DOI:10.1161/atv81.7.3.3593075.

(251)

Davies, P. F.; Robotewskyj, A.; Griem, M. L. Quantitative Studies of Endothelial Cell Adhesion. Directional Remodeling of Focal Adhesion Sites in Response to Flow Forces. J. Clin. Invest. 1994, 93 (5), 2031–2038. DOI:10.1172/JCI117197.

(252)

Yoshino, D.; Sakamoto, N.; Sato, M. Fluid Shear Stress Combined with Shear Stress Spatial Gradients Regulates Vascular Endothelial Morphology. Integr. Biol. 2017, 9 (7), 584–594. DOI:10.1039/C7IB00065K.

(253)

Yoshino, D.; Sakamoto, N.; Takahashi, K.; Inoue, E.; Sato, M. Development of Novel Flow Chamber to Study Endothelial Cell Morphology: Effects of Shear Flow with Uniform Spatial Gradient on Distribution of Focal Adhesion. J. Biomech. Sci. Eng. 2013, 8 (3), 233–243. DOI:10.1299/jbse.8.233.

(254)

Moretti, M.; Prina-Mello, A.; Reid, A. J.; Barron, V.; Prendergast, P. J. Endothelial Cell Alignment on Cyclically-Stretched Silicone Surfaces. J. Mater. Sci. Mater. Med. 2004, 15 (10), 1159–1164.



DOI:10.1023/B:JMSM.0000046400.18607.72. (255)

Yoshigi, M.; Clark, E. B.; Yost, H. J. Quantification of Stretch-Induced Cytoskeletal Remodeling in Vascular Endothelial Cells by Image Processing. Cytometry 2003, 55A (2), 109–118. DOI:10.1002/cyto.a.10076.

(256)

Barron, V.; Brougham, C.; Coghlan, K.; McLucas, E.; O’Mahoney, D.; Stenson-Cox, C.; McHugh, P. E. The Effect of Physiological Cyclic Stretch on the Cell Morphology, Cell Orientation and Protein Expression of Endothelial Cells. J. Mater. Sci. Mater. Med. 2007, 18 (10), 1973–1981. DOI:10.1007/s10856-007-31253.

(257)

Iwayoshi, S.; Furukawa, K.; Ushida, T. Continuous Visualization of Morphological Changes in Endothelial Cells in Response to Cyclic Stretch. JSME Int. J. Ser. C 2006, 49 (2), 545–555. DOI:10.1299/jsmec.49.545.

(258)

Ives, C. L.; Eskin, S. G.; McIntire, L. V. Mechanical Effects on Endothelial Cell Morphology: In Vitro Assessment. Vitr. Cell. Dev. Biol. 1986, 22 (9), 500–507. DOI:10.1007/BF02621134.

(259)

Haghighipour, N.; Tafazzoli-Shadpour, M.; Shokrgozar, M. A.; Amini, S.; Amanzadeh, A.; Khorasani, M. T. Topological Remodeling of Cultured Endothelial Cells by Characterized Cyclic Strains. Mol. Cell. Biomech. 2007, 4 (4), 189–199. DOI:10.3970/mcb.2007.004.189.

(260)

Hatami, J.; Tafazzoli-Shadpour, M.; Haghighipour, N.; Shokrgozar, M. A.; Janmaleki, M. Influence of Cyclic Stretch on Mechanical Properties of Endothelial Cells. Exp. Mech. 2013, 53 (8), 1291–1298. DOI:10.1007/s11340-013-9744-3.

(261)

von Offenberg, S. N.; Cummins, P. M.; Birney, Y. A.; Cullen, J. P.; Redmond, E. M.; Cahill, P. A. Cyclic Strain-Mediated Regulation of Endothelial Matrix Metalloproteinase-2 Expression and Activity. Cardiovasc. Res. 2004, 63 (4), 625–634. DOI:10.1016/j.cardiores.2004.05.008.

(262)

Wang, B.-W.; Chang, H.; Lin, S.; Kuan, P.; Shyu, K.-G. Induction of Matrix Metalloproteinases-14 and -2 by Cyclical Mechanical Stretch Is Mediated by Tumor Necrosis Factor-Alpha in Cultured Human Umbilical Vein Endothelial Cells. Cardiovasc. Res. 2003, 59 (2), 460–469. DOI:10.1016/S0008-6363(03)00428-0.

(263)

Yung, Y. C.; Chae, J.; Buehler, M. J.; Hunter, C. P.; Mooney, D. J. Cyclic Tensile Strain Triggers a Sequence of Autocrine and Paracrine Signaling to Regulate Angiogenic Sprouting in Human Vascular Cells. Proc. Natl. Acad. Sci. 2009, 106 (36), 15279–15284. DOI:10.1073/pnas.0905891106.

(264)

Wilkins, J. R.; Pike, D. B.; Gibson, C. C.; Kubota, A.; Shiu, Y.-T. Differential Effects of Cyclic Stretch on


Page 76 of 85

Page 77 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


BFGF- and VEGF-Induced Sprouting Angiogenesis. Biotechnol. Prog. 2014, 30 (4), 879–888. DOI:10.1002/btpr.1883. (265)

Bocci, G.; Fasciani, A.; Danesi, R.; Viacava, P.; Genazzani, A. R.; Del Tacca, M. In-Vitro Evidence of Autocrine Secretion of Vascular Endothelial Growth Factor by Endothelial Cells from Human Placental Blood Vessels. Mol. Hum. Reprod. 2001, 7 (8), 771–777. DOI:10.1093/molehr/7.8.771.

(266)

Tian, Y.; Gawlak, G.; O’Donnell, J. J.; Birukova, A. A.; Birukov, K. G. Activation of Vascular Endothelial Growth Factor (VEGF) Receptor 2 Mediates Endothelial Permeability Caused by Cyclic Stretch. J. Biol. Chem. 2016, 291 (19), 10032–10045. DOI:10.1074/jbc.M115.690487.

(267)

Toda, M.; Yamamoto, K.; Shimizu, N.; Obi, S.; Kumagaya, S.; Igarashi, T.; Kamiya, A.; Ando, J. Differential Gene Responses in Endothelial Cells Exposed to a Combination of Shear Stress and Cyclic Stretch. J. Biotechnol. 2008, 133 (2), 239–244. DOI:10.1016/j.jbiotec.2007.08.009.

(268)

Fisslthaler, B.; Popp, R.; Michaelis, U. R.; Kiss, L.; Fleming, I.; Busse, R. Cyclic Stretch Enhances the Expression and Activity of Coronary Endothelium-Derived Hyperpolarizing Factor Synthase. Hypertension 2001, 38 (6), 1427–1432. DOI:10.1161/hy1201.096532.

(269)

Takeda, H.; Komori, K.; Nishikimi, N.; Nimura, Y.; Sokabe, M.; Naruse, K. Bi-Phasic Activation of ENOS in Response to Uni-Axial Cyclic Stretch Is Mediated by Differential Mechanisms in BAECs. Life Sci. 2006, 79 (3), 233–239. DOI:10.1016/j.lfs.2005.12.051.

(270)

Carosi, J. A.; Eskin, S. G.; McIntire, L. V. Cyclical Strain Effects on Production of Vasoactive Materials in Cultured Endothelial Cells. J. Cell. Physiol. 1992, 151 (1), 29–36. DOI:10.1002/jcp.1041510106.

(271)

Carosi, J. A.; McIntire, L. V.; Eskin, S. G. Modulation of Secretion of Vasoactive Materials from Human and Bovine Endothelial Cells by Cyclic Strain. Biotechnol. Bioeng. 1994, 43 (7), 615–621. DOI:10.1002/bit.260430711.

(272)

Ali, M. H.; Pearlstein, D. P.; Mathieu, C. E.; Schumacker, P. T. Mitochondrial Requirement for Endothelial Responses to Cyclic Strain: Implications for Mechanotransduction. Am. J. Physiol. Cell. Mol. Physiol. 2004, 287 (3), L486–L496. DOI:10.1152/ajplung.00389.2003.

(273)

Goettsch, C.; Goettsch, W.; Arsov, A.; Hofbauer, L. C.; Bornstein, S. R.; Morawietz, H. Long-Term Cyclic Strain Downregulates Endothelial Nox4. Antioxid. Redox Signal. 2009, 11 (10), 2385–2397. DOI:10.1089/ars.2009.2561.



(274)

Sugawara, E.; Nikaido, H. Properties of AdeABC and AdeIJK Efflux Systems of Acinetobacter Baumannii Compared with Those of the AcrAB-TolC System of Escherichia Coli. Antimicrob. Agents Chemother. 2014, 58 (12), 7250–7257. DOI:10.1128/AAC.03728-14.

(275)

Li, W.; Sumpio, B. E. Strain-Induced Vascular Endothelial Cell Proliferation Requires PI3K-Dependent MTOR-4E-BP1 Signal Pathway. Am. J. Physiol. Circ. Physiol. 2005, 288 (4), H1591–H1597. DOI:10.1152/ajpheart.00382.2004.

(276)

Hurley, N. E.; Schildmeyer, L. A.; Bosworth, K. A.; Sakurai, Y.; Eskin, S. G.; Hurley, L. H.; McIntire, L. V. Modulating the Functional Contributions of C-Myc to the Human Endothelial Cell Cyclic Strain Response. J. Vasc. Res. 2010, 47 (1), 80–90. DOI:10.1159/000235928.

(277)

Schwartz, E. A.; Gerritsen, M. E.; Bizios, R. Effects of Hydrostatic Pressure on Endothelial Cells. In Mechanical Forces and the Endothelium; Lelkes, P. I., Ed.; CRC Press, 1999; pp 275–290.

(278)

Sumpio, B. E.; Widmann, M. D.; Ricotta, J.; Awolesi, M. A.; Watase, M. Increased Ambient Pressure Stimulates Proliferation and Morphologic Changes in Cultured Endothelial Cells. J. Cell. Physiol. 1994, 158 (1), 133–139. DOI:10.1002/jcp.1041580117.

(279)

Acevedo, A. D.; Bowser, S. S.; Gerritsen, M. E.; Bizios, R. Morphological and Proliferative Responses of Endothelial Cells to Hydrostatic Pressure: Role of Fibroblast Growth Factor. J. Cell. Physiol. 1993, 157 (3), 603–614. DOI:10.1002/jcp.1041570321.

(280)

Sugaya, Y.; Sakamoto, N.; Ohashi, T.; Sato, M. Elongation and Random Orientation of Bovine Endothelial Cells in Response to Hydrostatic Pressure: Comparison with Response to Shear Stress. JSME Int. J. Ser. C 2003, 46 (4), 1248–1255. DOI:10.1299/jsmec.46.1248.

(281)

Schwartz, E. A.; Bizios, R.; Medow, M. S.; Gerritsen, M. E. Exposure of Human Vascular Endothelial Cells to Sustained Hydrostatic Pressure Stimulates Proliferation. Involvement of the AlphaV Integrins. Circ. Res. 1999, 84 (3), 315–322. DOI:10.1161/res.84.3.315.

(282)

Ohashi, T.; Sugaya, Y.; Sakamoto, N.; Sato, M. Hydrostatic Pressure Influences Morphology and Expression of VE-Cadherin of Vascular Endothelial Cells. J. Biomech. 2007, 40 (11), 2399–2405. DOI:10.1016/j.jbiomech.2006.11.023.

(283)

Tokunaga, O.; Fan, J. L.; Watanabe, T. Atherosclerosis and Endothelium. Part II. Properties of Aortic Endothelial and Smooth Muscle Cells Cultured at Various Ambient Pressures. Pathol. Int. 1989, 39 (6),


Page 78 of 85

Page 79 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


356–362. DOI:10.1111/j.1440-1827.1989.tb02447.x. (284)

Shin, H. Y.; Gerritsen, M. E.; Bizios, R. Regulation of Endothelial Cell Proliferation and Apoptosis by Cyclic Pressure. Ann. Biomed. Eng. 2002, 30 (3), 297–304. DOI:10.1114/1.1458595.

(285)

Shin, H. Y.; Schwartz, E. A.; Bizios, R.; Gerritsen, M. E. Receptor-Mediated Basic Fibroblast Growth Factor Signaling Regulates Cyclic Pressure–Induced Human Endothelial Cell Proliferation. Endothelium 2004, 11 (5–6), 285–291. DOI:10.1080/10623320490904205.

(286)

Shin, H. Y.; Smith, M. L.; Toy, K. J.; Williams, P. M.; Bizios, R.; Gerritsen, M. E. VEGF-C Mediates Cyclic Pressure-Induced Endothelial Cell Proliferation. Physiol. Genomics 2002, 11 (3), 245–251. DOI:10.1152/physiolgenomics.00068.2002.

(287)

Hishikawa, K.; Nakaki, T.; Marumo, T.; Suzuki, H.; Kato, R.; Saruta, T. Pressure Enhances Endothelin-1 Release From Cultured Human Endothelial Cells. Hypertension 1995, 25 (3), 449–452. DOI:10.1161/01.HYP.25.3.449.

(288)

Li, L.; Yang, Y.; Shi, X.; Wu, H.; Chen, H.; Liu, J. A Microfluidic System for the Study of the Response of Endothelial Cells under Pressure. Microfluid. Nanofluidics 2014, 16 (6), 1089–1096. DOI:10.1007/s10404013-1275-9.

(289)

Vozzi, F.; Bianchi, F.; Ahluwalia, A.; Domenici, C. Hydrostatic Pressure and Shear Stress Affect Endothelin-1 and Nitric Oxide Release by Endothelial Cells in Bioreactors. Biotechnol. J. 2014, 9 (1), 146– 154. DOI:10.1002/biot.201300016.

(290)

Yang, J. J.; Chen, Y. M.; Kurokawa, T.; Gong, J. P.; Onodera, S.; Yasuda, K. Gene Expression, Glycocalyx Assay, and Surface Properties of Human Endothelial Cells Cultured on Hydrogel Matrix with Sulfonic Moiety: Effect of Elasticity of Hydrogel. J. Biomed. Mater. Res. A 2010, 95 (2), 531–542. DOI:10.1002/jbm.a.32875.

(291)

Yeh, Y.-T.; Hur, S. S.; Chang, J.; Wang, K.-C.; Chiu, J.-J.; Li, Y.-S.; Chien, S. Matrix Stiffness Regulates Endothelial Cell Proliferation through Septin 9. PLoS One 2012, 7 (10), e46889. DOI:10.1371/journal.pone.0046889.

(292)

Chen, J.; Hu, M.; Zhang, H.; Li, B.; Chang, H.; Ren, K.; Wang, Y.; Ji, J. Improved Antithrombotic Function of Oriented Endothelial Cell Monolayer on Microgrooves. ACS Biomater. Sci. Eng. 2017, acsbiomaterials.7b00496. DOI:10.1021/acsbiomaterials.7b00496.



(293)

Lamichhane, S.; Anderson, J. A.; Remund, T.; Sun, H.; Larson, M. K.; Kelly, P.; Mani, G. Responses of Endothelial Cells, Smooth Muscle Cells, and Platelets Dependent on the Surface Topography of Polytetrafluoroethylene. J. Biomed. Mater. Res. Part A 2016, 104 (9), 2291–2304. DOI:10.1002/jbm.a.35763.

(294)

McKee, C. T.; Wood, J. A.; Ly, I.; Russell, P.; Murphy, C. J. The Influence of a Biologically Relevant Substratum Topography on Human Aortic and Umbilical Vein Endothelial Cells. Biophys. J. 2012, 102 (5), 1224–1233. DOI:10.1016/j.bpj.2012.01.053.

(295)

Cutiongco, M. F. A.; Chua, B. M. X.; Neo, D. J. H.; Rizwan, M.; Yim, E. K. F. Functional Differences between Healthy and Diabetic Endothelial Cells on Topographical Cues. Biomaterials 2018, 153, 70–84. DOI:10.1016/j.biomaterials.2017.10.037.

(296)

Moore, J. E.; Bürki, E.; Suciu, A.; Zhao, S.; Burnier, M.; Brunner, H. R.; Meister, J.-J. A Device for Subjecting Vascular Endothelial Cells to Both Fluid Shear Stress and Circumferential Cyclic Stretch. Ann. Biomed. Eng. 1994, 22 (4), 416–422. DOI:10.1007/BF02368248.

(297)

Zheng, W.; Jiang, B.; Wang, D.; Zhang, W.; Wang, Z.; Jiang, X. A Microfluidic Flow-Stretch Chip for Investigating Blood Vessel Biomechanics. Lab Chip 2012, 12 (18), 3441. DOI:10.1039/c2lc40173h.

(298)

Owatverot, T. B.; Oswald, S. J.; Chen, Y.; Wille, J. J.; Yin, F. C. P. Effect of Combined Cyclic Stretch and Fluid Shear Stress on Endothelial Cell Morphological Responses. J. Biomech. Eng. 2005, 127 (3), 374–382. DOI:10.1115/1.1894180.

(299)

Sinha, R.; Le Gac, S.; Verdonschot, N.; van den Berg, A.; Koopman, B.; Rouwkema, J. Endothelial Cell Alignment as a Result of Anisotropic Strain and Flow Induced Shear Stress Combinations. Sci. Rep. 2016, 6 (1), 29510. DOI:10.1038/srep29510.

(300)

Qiu, Y.; Tarbell, J. M. Interaction between Wall Shear Stress and Circumferential Strain Affects Endothelial Cell Biochemical Production. J. Vasc. Res. 2000, 37 (3), 147–157. DOI:10.1159/000025726.

(301)

Ziegler, T.; Silacci, P.; Harrison, V. J.; Hayoz, D. Nitric Oxide Synthase Expression in Endothelial Cells Exposed to Mechanical Forces. Hypertension 1998, 32 (2), 351–355. DOI:10.1161/hyp.32.2.351.

(302)

Dancu, M. B.; Berardi, D. E.; Vanden Heuvel, J. P.; Tarbell, J. M. Asynchronous Shear Stress and Circumferential Strain Reduces Endothelial NO Synthase and Cyclooxygenase-2 but Induces Endothelin-1 Gene Expression in Endothelial Cells. Arterioscler. Thromb. Vasc. Biol. 2004, 24 (11), 2088–2094.


Page 80 of 85

Page 81 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


DOI:10.1161/01.ATV.0000143855.85343.0e. (303)

Uttayarat, P.; Chen, M.; Li, M.; Allen, F. D.; Composto, R. J.; Lelkes, P. I. Microtopography and Flow Modulate the Direction of Endothelial Cell Migration. Am. J. Physiol. Circ. Physiol. 2008, 294 (2), H1027– H1035. DOI:10.1152/ajpheart.00816.2007.

(304)

Morgan, J. T.; Wood, J. A.; Shah, N. M.; Hughbanks, M. L.; Russell, P.; Barakat, A. I.; Murphy, C. J. Integration of Basal Topographic Cues and Apical Shear Stress in Vascular Endothelial Cells. Biomaterials 2012, 33 (16), 4126–4135. DOI:10.1016/j.biomaterials.2012.02.047.

(305)

Franco, D.; Milde, F.; Klingauf, M.; Orsenigo, F.; Dejana, E.; Poulikakos, D.; Cecchini, M.; Koumoutsakos, P.; Ferrari, A.; Kurtcuoglu, V. Accelerated Endothelial Wound Healing on Microstructured Substrates under Flow. Biomaterials 2013, 34 (5), 1488–1497. DOI:10.1016/j.biomaterials.2012.10.007.

(306)

Robotti, F.; Franco, D.; Bänninger, L.; Wyler, J.; Starck, C. T.; Falk, V.; Poulikakos, D.; Ferrari, A. The Influence of Surface Micro-Structure on Endothelialization under Supraphysiological Wall Shear Stress. Biomaterials 2014, 35 (30), 8479–8486. DOI:10.1016/j.biomaterials.2014.06.046.

(307)

Hu, J.; Hardy, C.; Chen, C.-M.; Yang, S.; Voloshin, A. S.; Liu, Y. Enhanced Cell Adhesion and Alignment on Micro-Wavy Patterned Surfaces. PLoS One 2014, 9 (8), e104502. DOI:10.1371/journal.pone.0104502.

(308)

Whited, B. M.; Rylander, M. N. The Influence of Electrospun Scaffold Topography on Endothelial Cell Morphology, Alignment, and Adhesion in Response to Fluid Flow. Biotechnol. Bioeng. 2014, 111 (1), 184– 195. DOI:10.1002/bit.24995.

(309)

Kohn, J. C.; Zhou, D. W.; Bordeleau, F.; Zhou, A. L.; Mason, B. N.; Mitchell, M. J.; King, M. R.; ReinhartKing, C. A. Cooperative Effects of Matrix Stiffness and Fluid Shear Stress on Endothelial Cell Behavior. Biophys. J. 2015, 108 (3), 471–478. DOI:10.1016/j.bpj.2014.12.023.

(310)

Dan, A.; Huang, R. B.; Leckband, D. E. Dynamic Imaging Reveals Coordinate Effects of Cyclic Stretch and Substrate Stiffness on Endothelial Integrity. Ann. Biomed. Eng. 2016, 44 (12), 3655–3667. DOI:10.1007/s10439-016-1677-4.

(311)

Benbrahim, A.; L’Italien, G. J.; Kwolek, C. J.; Petersen, M. J.; Milinazzo, B.; Gertler, J. P.; Abbott, W. M.; Orkin, R. W. Characteristics of Vascular Wall Cells Subjected to Dynamic Cyclic Strain and Fluid Shear Conditions in Vitro. J. Surg. Res. 1996, 65 (2), 119–127. DOI:10.1006/jsre.1996.0353.

(312)

Benbrahim, A.; L’Italien, G. J.; Milinazzo, B. B.; Warnock, D. F.; Dhara, S.; Gertler, J. P.; Orkin, R. W.;



Abbott, W. M. A Compliant Tubular Device to Study the Influences of Wall Strain and Fluid Shear Stress on Cells of the Vascular Wall. J. Vasc. Surg. 1994, 20 (2), 184–194. DOI:10.1016/0741-5214(94)90005-1. (313)

Peng, X.; Recchia, F. A.; Byrne, B. J.; Wittstein, I. S.; Ziegelstein, R. C.; Kass, D. A. In Vitro System to Study Realistic Pulsatile Flow and Stretch Signaling in Cultured Vascular Cells. Am. J. Physiol. Cell Physiol. 2000, 279 (3), C797-805. DOI:10.1152/ajpcell.2000.279.3.C797.

(314)

Estrada, R.; Giridharan, G. A.; Nguyen, M.-D.; Roussel, T. J.; Shakeri, M.; Parichehreh, V.; Prabhu, S. D.; Sethu, P. Endothelial Cell Culture Model for Replication of Physiological Profiles of Pressure, Flow, Stretch, and Shear Stress in Vitro. Anal. Chem. 2011, 83 (8), 3170–3177. DOI:10.1021/ac2002998.

(315)

Estrada, R.; Giridharan, G. A.; Nguyen, M.-D.; Prabhu, S. D.; Sethu, P. Microfluidic Endothelial Cell Culture Model to Replicate Disturbed Flow Conditions Seen in Atherosclerosis Susceptible Regions. Biomicrofluidics 2011, 5 (3), 032006. DOI:10.1063/1.3608137.

(316)

Casey, P. J.; Dattilo, J. B.; Dai, G.; Albert, J. A.; Tsukurov, O. I.; Orkin, R. W.; Gertler, J. P.; Abbott, W. M. The Effect of Combined Arterial Hemodynamics on Saphenous Venous Endothelial Nitric Oxide Production. J. Vasc. Surg. 2001, 33 (6), 1199–1205. DOI:10.1067/mva.2001.115571.

(317)

Tsukurov, O. I.; Kwolek, C. J.; L’Italien, G. J.; Benbrahim, A.; Milinazzo, B. B.; Conroy, N. E.; Gertler, J. P.; Orkin, R. W.; Abbott, W. M. The Response of Adult Human Saphenous Vein Endothelial Cells to Combined Pressurized Pulsatile Flow and Cyclic Strain, In Vitro. Ann. Vasc. Surg. 2000, 14 (3), 260–267. DOI:10.1007/s100169910044.

(318)

Tarbell, J. M. The Interaction between Fluid-Wall Shear Stress and Solid Circumferential Strain Affects Endothelial Cell Mechanobiology. In Cellular Mechanotransduction; Mofrad, M. R. K., Kamm, R. D., Eds.; Cambridge University Press: Cambridge; pp 360–376. DOI:10.1017/CBO9781139195874.016.

(319)

Meza, D.; Abejar, L.; Rubenstein, D. A.; Yin, W. A Shearing-Stretching Device That Can Apply Physiological Fluid Shear Stress and Cyclic Stretch Concurrently to Endothelial Cells. J. Biomech. Eng. 2016, 138 (3), 031007. DOI:10.1115/1.4032550.

(320)

Regan, E. R.; Aird, W. C. Dynamical Systems Approach to Endothelial Heterogeneity. Circ. Res. 2012, 111 (1), 110–130. DOI:10.1161/CIRCRESAHA.111.261701.

(321)

Jiang, Y.-Z.; Manduchi, E.; Jimenez, J. M.; Davies, P. F. Endothelial Epigenetics in Biomechanical Stress: Disturbed Flow-Mediated Epigenomic Plasticity In Vivo and In Vitro. Arterioscler. Thromb. Vasc. Biol.


Page 82 of 85

Page 83 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


2015, 35 (6), 1317–1326. DOI:10.1161/ATVBAHA.115.303427. (322)

Montgomery, D. C. Design and Analysis of Experiments, 8th ed.; John Wiley & Sons, Inc.: Hoboken, NJ, 2012.

(323)

Haase, K.; Kamm, R. D. Advances in On-Chip Vascularization. Regen. Med. 2017, 12 (3), 285–302. DOI:10.2217/rme-2016-0152.

(324)

Moore, E. M.; Ying, G.; West, J. L. Macrophages Influence Vessel Formation in 3D Bioactive Hydrogels. Adv. Biosyst. 2017, 1 (3), 1600021. DOI:10.1002/adbi.201600021.

(325)

Rivron, N. C.; Vrij, E. J.; Rouwkema, J.; Le Gac, S.; van den Berg, A.; Truckenmuller, R. K.; van Blitterswijk, C. A. Tissue Deformation Spatially Modulates VEGF Signaling and Angiogenesis. Proc. Natl. Acad. Sci. 2012, 109 (18), 6886–6891. DOI:10.1073/pnas.1201626109.

(326)

Ingber, D. E. Mechanical Signaling and the Cellular Response to Extracellular Matrix in Angiogenesis and Cardiovascular Physiology. Circ. Res. 2002, 91 (10), 877–887. DOI:10.1161/01.RES.0000039537.73816.E5.

(327)

Kannan, R. Y.; Salacinski, H. J.; Sales, K.; Butler, P.; Seifalian, A. M. The Roles of Tissue Engineering and Vascularisation in the Development of Micro-Vascular Networks: A Review. Biomaterials 2005, 26 (14), 1857–1875. DOI:10.1016/j.biomaterials.2004.07.006.

(328)

Song, J. W.; Munn, L. L. Fluid Forces Control Endothelial Sprouting. Proc. Natl. Acad. Sci. 2011, 108 (37), 15342–15347. DOI:10.1073/pnas.1105316108.

(329)

Zheng, Y.; Chen, J.; Craven, M.; Choi, N. W.; Totorica, S.; Diaz-Santana, A.; Kermani, P.; Hempstead, B.; Fischbach-Teschl, C.; Lopez, J. A.; et al. In Vitro Microvessels for the Study of Angiogenesis and Thrombosis. Proc. Natl. Acad. Sci. 2012, 109 (24), 9342–9347. DOI:10.1073/pnas.1201240109.

(330)

Yeon, J. H.; Ryu, H. R.; Chung, M.; Hu, Q. P.; Jeon, N. L. In Vitro Formation and Characterization of a Perfusable Three-Dimensional Tubular Capillary Network in Microfluidic Devices. Lab Chip 2012, 12 (16), 2815. DOI:10.1039/c2lc40131b.

(331)

Hosseini, Y.; . Agah, M.; . Verbridge, S. S. Endothelial Cell Sensing, Restructuring, and Invasion in Collagen Hydrogel Structures. Integr. Biol. 2015, 7 (11), 1432–1441. DOI:10.1039/C5IB00207A.

(332)

Aizawa, Y.; Wylie, R.; Shoichet, M. Endothelial Cell Guidance in 3D Patterned Scaffolds. Adv. Mater. 2010, 22 (43), 4831–4835. DOI:10.1002/adma.201001855.



(333)

Rosenfeld, D.; Landau, S.; Shandalov, Y.; Raindel, N.; Freiman, A.; Shor, E.; Blinder, Y.; Vandenburgh, H. H.; Mooney, D. J.; Levenberg, S. Morphogenesis of 3D Vascular Networks Is Regulated by Tensile Forces. Proc. Natl. Acad. Sci. 2016, 113 (12), 3215–3220. DOI:10.1073/pnas.1522273113.

(334)

Nsiah, B. A.; Moore, E. M.; Roudsari, L. C.; Virdone, N. K.; West, J. L. Angiogenesis in Hydrogel Biomaterials. In Biosynthetic Polymers for Medical Applications; Elsevier, 2016; pp 189–203. DOI:10.1016/B978-1-78242-105-4.00008-0.

(335)

Dao Thi, M.-U.; Trocmé, C.; Montmasson, M.-P.; Fanchon, E.; Toussaint, B.; Tracqui, P. Investigating Metalloproteinases MMP-2 and MMP-9 Mechanosensitivity to Feedback Loops Involved in the Regulation of In Vitro Angiogenesis by Endogenous Mechanical Stresses. Acta Biotheor. 2012, 60 (1–2), 21–40. DOI:10.1007/s10441-012-9147-3.

(336)

Philpott, A. C.; Southern, D. A.; Clement, F. M.; Galbraith, P. D.; Traboulsi, M.; Knudtson, M. L.; Ghali, W. A. Long-Term Outcomes of Patients Receiving Drug-Eluting Stents. Can. Med. Assoc. J. 2009, 180 (2), 167–174. DOI:10.1503/cmaj.080050.

(337)

Bønaa, K. H.; Mannsverk, J.; Wiseth, R.; Aaberge, L.; Myreng, Y.; Nygård, O.; Nilsen, D. W.; Kløw, N.-E.; Uchto, M.; Trovik, T.; et al. Drug-Eluting or Bare-Metal Stents for Coronary Artery Disease. N. Engl. J. Med. 2016, 375 (13), 1242–1252. DOI:10.1056/NEJMoa1607991.

(338)

Singh, C.; Wong, C.; Wang, X. Medical Textiles as Vascular Implants and Their Success to Mimic Natural Arteries. J. Funct. Biomater. 2015, 6 (3), 500–525. DOI:10.3390/jfb6030500.

(339)

Silverthorn, D. U. Human Physiology: An Integrated Approach, 4th ed.; Pearson/Benjamin Cummings, 2007.

(340)

Pashneh-Tala, S.; MacNeil, S.; Claeyssens, F. The Tissue-Engineered Vascular Graft—Past, Present, and Future. Tissue Eng. Part B Rev. 2015, ten.teb.2015.0100. DOI:10.1089/ten.teb.2015.0100.


Page 84 of 85

Page 85 of 85 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


For Table of Contents Use Only

Title: Vascular endothelial cell behavior in complex mechanical microenvironments Authors: Bryan D. James and Josephine Allen


Vascular Endothelial Cell Behavior in Complex Mechanical

Recommend Documents