Cell Mechanics: Integrating Cell Responses to Mechanical ... · ANRV317-BE09-01 ARI 7 June 2007...

ANRV317-BE09-01 ARI 7 June 2007 15:9

Cell Mechanics: IntegratingCell Responses toMechanical StimuliPaul A. Janmey1 and Christopher A. McCulloch2

1Departments of Physiology, Physics, and Bioengineering, Institute for Medicineand Engineering, University of Pennsylvania, Philadelphia, Pennsylvania 19104;email: [email protected] Group in Matrix Dynamics, Fitzgerald Building, University of Toronto,Toronto, Ontario, M5S 3E2 Canada; email: [email protected]

Annu. Rev. Biomed. Eng. 2007. 9:1–34

First published online as a Review in Advance onApril 26, 2007

The Annual Review of Biomedical Engineering isonline at bioeng.annualreviews.org

This article’s doi:10.1146/annurev.bioeng.9.060906.151927

Copyright c© 2007 by Annual Reviews.All rights reserved

1523-9829/07/0815-0001$20.00

AbstractForces are increasingly recognized as major regulators of cell struc-ture and function, and the mechanical properties of cells are essentialto the mechanisms by which cells sense forces, transmit them to thecell interior or to other cells, and transduce them into chemical sig-nals that impact a spectrum of cellular responses. Comparison ofthe mechanical properties of intact cells with those of the purifiedcytoskeletal biopolymers that are thought to dominate their elas-ticity reveal the extent to which the studies of purified systems canaccount for the mechanical properties of the much more heteroge-neous and complex cell. This review summarizes selected aspects ofcurrent work on cell mechanics with an emphasis on the structuresthat are activated in cell-cell contacts, that regulate ion flow acrossthe plasma membrane, and that may sense fluid flow that produceslow levels of shear stress.

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Contents

INTRODUCTION. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2MEASUREMENTS OF CELL STIFFNESS IN VITRO . . . . . . . . . . . . . . 3MODELS FOR CELL MECHANICS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

Viscoelastic Gels of Cytoskeletal Polymers . . . . . . . . . . . . . . . . . . . . . . . . . . . 5Colloidal, Glassy, and Foam Models for Cell Mechanics . . . . . . . . . . . . . . 7Time Dependence of Cell Elasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

IMPLICATIONS OF CELL MECHANICAL PROPERTIES FORTHE MECHANISMS BY WHICH FORCES ARE SENSED . . . . . . . 8The Extracellular Matrix in Cell Mechanics . . . . . . . . . . . . . . . . . . . . . . . . . . 9Attachment Receptors in Force Transmission and Sensing . . . . . . . . . . . . 10Cadherins and Mechanobiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11Adhesion Systems and the Cytoskeleton in Regulation

of Ion Channels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12Force Transmission to Intracellular Organelles . . . . . . . . . . . . . . . . . . . . . . . 13

HOW ARE FORCES EXERTED AT THE CELL SURFACETRANSMITTED THROUGH THE CELL? . . . . . . . . . . . . . . . . . . . . . . 14Theoretical Predictions of the Stress Field in a Model Cell . . . . . . . . . . . 15Experimental Measurements of Intracellular Strains Caused

by Extracellular Forces . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16SENSING AND RESPONDING TO SHEAR STRESS CAUSED BY

FLUID FLOW . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18Primary Cilium. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19Bending of the Primary Cilium by Fluid Flow. . . . . . . . . . . . . . . . . . . . . . . . 20

CONCLUSIONS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22

INTRODUCTION

The mechanical properties of cells have attracted interest from the very beginning ofcell biologic studies. Gel-sol transitions were recognized in the cytoplasm of crawlingamoeboid cells in the eighteenth century (1) and flow within plant cells providedsome of the first quantitative data describing non-Newtonian viscosity (reviewed inReference 2). In the 1920s microscopic magnetic particles were manipulated withinlive cells to obtain quantitative measurements of elastic and viscous parameters bymicrorheology (3, 4). In part, the focus on cell mechanics has been motivated byefforts to define how cells perform mechanical work as they move (5); however,recently, increased interest in cell mechanics has been generated by demonstrationsthat the mechanical features of the extracellular environment and application of forcesto cells trigger cellular responses that are essential for many aspects of cell structureand function (reviewed in References 6, 7). This review focuses on examples of howcells respond to specific types of mechanical stresses, how the mechanical propertiesof cells might relate to the rheological properties of synthetic or model systems, and

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AFM: atomic forcemicroscopy

how the mechanical features of cytoskeletal elements provide the potential to detectand integrate physical stimuli.

In contrast to the level of understanding of how specific chemicals trigger andtransduce signals within cells, studies of how cells sense force and how they respondto different levels, durations, and directions of force are much less common, but haverecently attracted increasing attention from biologists and bioengineers. Definingspecific structures and mechanisms by which forces—both external and internallygenerated—are sensed by cells and how this stimulus leads to specific responses islikely to help explain the complex functions of cells and to design better materials forcell and tissue engineering and other applications in vivo.

MEASUREMENTS OF CELL STIFFNESS IN VITRO

The mechanical properties of the cell are essential to force responses because theydetermine the extent of deformation of the region of the cell where the force is applied,and therefore the range of molecular deformations that can be triggered. Estimatesof cellular stiffness derived from different types of microrheological methods covera wide range depending on the methods used, the type and extent of deformation,and the time that the deformation is applied, as shown in Table 1. The stiffnessmeasured for isolated cultured cells ranges from 0.1 kPa to an approximate upperlimit of near 40 kPa (8) for myocytes, similar to the range of stiffness measured formany soft tissues (9). The magnitudes of these moduli can help determine whichmolecular structures are responsible for cellular stiffness. For example, a reasonableestimate for the concentration of cytoskeletal proteins is 10 mg/ml actin, and tubulinand intermediate filament proteins have similar but slightly lower concentrations.In vitro studies show that cross-linked networks of actin at this concentration haveelastic moduli in the range of 10 to 5,000 Pa, as do intermediate filaments networks,but only when deformed to large strains (10, 14). Microtubules may also contributeto cellular stiffness if bound to the other filament systems, but most studies in whichmicrotubules are depolymerized show little effect of the microtubules directly on cellelasticity except when their depolymerization leads to rearrangement or disassemblyof the other filament types (11, 12).

The dominance of actin in determining the stiffness of cells is shown by one recentstudy using atomic force microscopy (AFM) to quantify stiffness from small indenta-tions of less than 200 nm at the dorsal surface of endothelial cells (13). These studiesreveal a heterogeneous population of local stiffness values that separated into two setswith elastic moduli of 5.6+/−3.5 kPa and 1.5+/−0.76 kPa depending on whetherthere were actin-based stress fibers underneath the AFM probe. Addition of cytocha-lasin to disassemble the actin network reduced the stiffness to 0.89+/−0.46 kPa andthe network became more homogeneous. Measurements such as these argue stronglyin support of a contribution of cytoskeletal networks to cell rheology that can be in-terpreted at least in part by the viscoelasticity of cross-linked gels made in vitro bythe purified cytoskeletal components. For example, 0.7 mg/ml actin gels cross-linkedby alpha actinin have elastic moduli on the order of 100 Pa (14), and similar rigiditiesare seen for actin cross-linked by filamin A (15, 16). Because the elastic modulus of

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Table 1 Comparison of elastic moduli measured for single cells in culture

Cell type Elastic modulus (kPa) Method ReferenceRat aortic smooth muscle 1.5–11 Elongation between plates (180)Endothelial 1.5–5.6 AFM (13)Aortic endothelialNormal/ cholesterol depleted

0.32/0.54 Microaspiration (181)

Endothelial 0.5 cytoplasm5 nucleus

Uniaxial compression (182)

Inner hair cell 0.3 AFM (183)Outer hair cell 2–3.7 AFM (184)Cardiac myocytes 35–42 AFM (8)Fibroblast 0.6–1.6 AFM (185)Fibroblast 1–10 (differential stretch modulus) Uniaxial stretching/compression (21)Bovine articular chondrocytes 1.1–8 Creep cytoindentation apparatus (186)Chondrocytes, Endothelial 0.5 Microaspiration (187)Neutrophils passive/activated 0.38/0.8 AFM (188)C2C12 myoblasts 2 Cell loading device

(global compression)(189)

Alveolar epithelial 0.1–0.2 Magnetic twisting cytometry (190)Epithelial normal/cancerous 10–13/0.4 – 1.4 AFM (191)Osteoblast 1–2 AFM (192)FibroblastsNormal/transformed

0.22/0.19; 0.42–0.48/1.0 Optical stretcher (193, 194)

Melanoma 0.3–2.0 frequency dependent Magnetic twisting rheometry (195)Kidney epithelialCell cortexCell interior

0.160.04

Magnetic twisting rheometryTracer diffusion

(44)

3T3 fibroblast before/aftershear flow

0.015/ 0.06 Tracer diffusion (196)

C2-7 myogenic 0.66 Uniaxial stretching rheometer (197)

networks formed by semiflexible polymers is shown by experiment (10) and theory(17) to scale approximately with the square of the polymer concentration, 10 mg/mlcross-linked actin networks are predicted to have elastic moduli on the order of 10 kPaeven at low strains and in the absence of other strengthening effects, such as interac-tions with other filaments or from prestress owing to internal contraction by motors.

The values of elastic moduli calculated from force-deformation studies of cellsare inherently difficult to interpret even when the experimental results are highlyreproducible and well controlled. Among the factors that complicate the analysis arethe facts that the structure of the cell interior is not homogeneous; the earliest studiesof cell mechanics noted that many crawling cells have a hierarchy of layers with aliquid core, surrounded by one or two layers of solid material that either trappedor excluded organelles, and a very thin highly elastic membrane (18). Therefore,the elastic modulus calculated from subcellular deformations is model dependent.


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Further complications arise from the fact that the composition of the cytoskeleton,the density of activated cross-linkers, and the presence of other cellular componentsvary spatially and as a function of time. An additional feature evident in many studiesis that the elastic material is inherently nonlinear, in that its resistance to deforma-tion depends on the extent of deformation, even without active remodeling of thenetwork. This nonlinear elastic response is a consequence of the physical propertiesof semiflexible polymers and creates the potential for cells to modify their stiffness byapplying internal deformations, often called prestress, by activation of motors. A finalcomplication is that the elastic resistance of cells is probably not entirely due to thethermal motions of polymers that produce the familiar entropic elasticity of rubbers,nor to enthalpic bending of stiff rods, which is analogous to bending of macroscopicbeams. For example, when actin filaments are not tightly cross-linked to each other,activation of myosin motors arranged in antiparallel dimers increases fluidity becausethe long interdigitated filaments slide past each other more facilely when propelledby the actomyosin power stroke than by thermal motions (19). The finding that manyapparently random intracellular motions are much longer than can be accounted forby Brownian motion suggests that the rheology of the cytoskeleton and the cytoplasmin general may be substantially different from that of its constituents when they havecome to equilibrium.

Given all of these complexities, it is perhaps surprising that the stiffness of manydifferent cell types, measured using a variety of methods that impose a limited rangeof strains for times on the order of seconds, does not vary more widely and thatsignificant differences between different cell types can be identified. This findingsuggests that cells may work to maintain a tightly controlled rigidity under specificconditions, and therefore, deviations from a normal range of stiffness might be used toidentify abnormal cells (20). Moreover, cells change their stiffness when subjected toexternal stress. A recent study systematically varied the force applied to fibroblasts heldadherent to two parallel plates and measured the stiffness in response to incrementaldeformation performed when the cell was held under tension. This study showed thatthe incremental elastic modulus can change by nearly an order of magnitude whenthe cell is held under tension (21). The tension can arise either from applicationof external force to elongate the cell or by generation of internal stress owing toactive intracellular processes that increase tension when the cell is kept at a constantheight. These results support earlier studies showing that internal tension, often calledprestress, can increase the stiffness of cells. The tension or deformation-dependentincrease in the elastic modulus is consistent with the nonlinear viscoelasticity ofcytoskeletal networks, and it may allow for significant changes in cell stiffness withoutchanges in the amount of polymerized proteins.

MODELS FOR CELL MECHANICS

Viscoelastic Gels of Cytoskeletal Polymers

In some cases, biological materials are sufficiently uniform, or their mechanical prop-erties are dominated by only a single structure, and they can be reasonably interpreted


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by existing models for synthetic materials. For example, the mechanics of blood clotsand some extracellular matrices are sufficiently dependent on the polymer networksthat form them, i.e., fibrin (22–25) and collagen (26), respectively, that their rhe-ology resembles that of gels of cross-linked polymers. Their relatively frequency-independent shear storage moduli and low mechanical loss, as well as the scalingof elastic moduli with polymer concentrations (10, 17), all resemble those of othersemiflexible polymer networks. The relatively large elastic moduli for low volumefractions and strain-stiffening behavior distinguish these biomaterials from hydrogelsof flexible polymers, but they are relatively well modeled by theories for semiflexiblechains (17, 27, 28). The similarity in rheology between some extracellular materi-als and synthetic polymer gels and the strong effect of actin depolymerizing agentson cell mechanics has motivated attempts to account for cell rheology using modelsdeveloped for polymer networks.

The rheology of soft biological materials depends on the assembly of long proteinfilaments into networks of different geometries. The specific geometries of these net-works are determined by the chemical and mechanical properties of the filaments, aswell as those of the cross-links that hold the filaments together. Despite their chem-ical and structural differences, the most abundant protein filaments of extracellularmatrices and intracellular cytoskeletons have the common property of being semiflex-ible filaments cross-linked into open meshworks. Whereas some biopolymers suchas elastin have elastic properties very similar to rubber-like materials (29, 30), gelsformed by stiffer filaments, such as fibrin, collagen, actin, and intermediate filaments,have distinct rheological properties (28, 31). For example, they are able to formelastic gels at very low volume fractions, less than 0.01% under optimal conditions,and become stiffer the more they are deformed, a property termed strain-stiffening(28).

These mechanical properties likely contribute to the biological functions of cy-toskeletal proteins in forming elastic networks with minimal protein production andnetworks with large mesh sizes that allow passage of large macromolecular assembliesand organelles. The rheological properties of the networks formed in vivo are likelyto differ in some aspects from those formed in vitro. In vivo, the production of bothfilaments and networks is tightly regulated by numerous assembly-promoting andcross-linking factors (32, 33) that produce active networks with specified geometriesfar from equilibrium (19, 34–36) and not yet capable of being produced from purifiedfactors in vitro. Nevertheless, some unusual properties of cross-linked semiflexiblepolymer networks have recently been observed in live cells, lending support to theconcept that the network properties of isolated biopolymers can provide some insightinto cell mechanics. On the other hand, studies of isotropic biopolymer networks canonly partly reflect the mechanical properties of an intact cell because whole-cell rhe-ology results from deformations of multiple layers within the cell, spanning the thinelastic membrane, the subcortical actin gel, a more granular gel-like inner layer, amore fluid layer in the cell interior, and the nucleus, which is generally much stifferthan the rest of the cell (18). How these multiple layers together determine cell rhe-ology remains difficult to relate to the filamentous and other structures found withineach region.


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Colloidal, Glassy, and Foam Models for Cell Mechanics

Most biological materials, including single cells, are not well defined by a singlesynthetic analog. In part, this results from their structural complexity and the fact thatmany are under internal tension (21, 37–41), are constantly remodeling, or are subjectto apparently random, but nonthermal, fluctuations that are rarely, if ever, found insynthetic materials (42). Experimental evidence that cells were at least partly solidor could change from solid to liquid form was clear well before the concept of largemacromolecules was accepted and well before cytoskeletal filaments were visualizedby electron microscopy. Early explanations for cell rheology were based on conceptsof colloid science (43), in which interactions between small particles suspended insolvent, in this case granular material within the cytoplasm, led to solid-like rheologythat allowed flow as the tightly packed particles subjected to forces moved past eachother. Many aspects of the time dependence of cell rheology resemble the rheology ofcolloids, foams, or soft glassy materials, and recent systematic studies of cell elasticityand viscosity over much larger ranges of time and frequency than were previouslypossible have motivated researchers to challenge models of cell mechanics analogousto polymer gels.

Many recent studies have reported that cells and some purified systems mimickingthe cytoskeleton have elastic moduli that follow a power law with a small fractionalexponent, often near 0.1 to 0.2 over several orders of magnitude in time (38, 39, 44–50). This behavior is inconsistent with polymer models containing a small numberof relaxation times, or a finite longest relaxation time, as expected for a polymer gel,but rather it implies a continuum of relaxation times consistent with the rheology ofimmobilized colloids or soft glasses. The evidence in favor of power law rheology ofsingle cells and other soft tissues is increasingly documented, although the precisemolecular nature of this behavior remains unexplained. A related finding from mi-croscopic measurement of the fluctuations of particles within the cytoplasm showsthat while these motions appear random, they are much too large to be accounted forby thermal fluctuations. Instead, these motions seem to result from random jostlingof the cytoskeleton, membranes, and other structures that are in contact with motorproteins that produce mechanical motions of various kinds using the energy of ATPhydrolysis to do work (42). In many cases, the resulting motions are unidirectionalfor long distances and easily distinguished from Brownian motion, but in other casessingle steps in random directions could produce the apparently random but activemovements within the cell, with a resulting spectrum of relaxation times that couldcontribute to the power law behavior of the overall rheology.

Time Dependence of Cell Elasticity

Whether the cell appears to behave as a glass, a gel, or something else may depend onthe magnitude of the deformation and the time over which the response is observed.The power-law relation of the cell’s elastic modulus need not imply that the polymerproperties of the cytoskeleton are unimportant or even dominant for the rheology ofthe cell. The basic model for polymer network elasticity assumes that the filamentsremain chemically identical during deformation and that the cross-links between


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them are rigid compared with the filaments themselves. Neither of these assumptionsis necessarily true for live cells, and the active movement of filaments and otherstructures by motor proteins further complicates network rheology. Recent studiesexamining the viscoelasticity of cells and complex cytoskeletal protein networks showthat the time-dependent rheology of these systems can be similar. The multiple factorsthat mediate cytoskeletal networks, such as the size, stability, geometry, and flexibilityof cross-linkers; active motions; and perhaps changes in filament structure; can allcontribute to rheology to produce the spectrum of relaxation times inferred from thetime-dependent measurements of cells (51).

In summary, very few biological tissues are simple enough to approximate by anysingle rheological model. As a result, most rheological studies have been phenomeno-logical, with either a finite number of elastic and viscous elements coupled in series orparallel to mimic the rheological behavior, or scaling exponents and limiting valuesto define how they may be fit by glass-like models (45). These phenomenologicalmodels have been essential in many bioengineering contexts to develop protective ortherapeutic strategies. However, developing mechanical models by which the proper-ties of specific molecular structures, motor-derived forces, and cell-cell or cell-matrixinteractions account for tissue mechanics remains an area of active investigation.

IMPLICATIONS OF CELL MECHANICAL PROPERTIES FORTHE MECHANISMS BY WHICH FORCES ARE SENSED

Mechanosensation requires by definition an element that responds to an applied force,without necessarily being triggered by binding a specific chemical stimulus. One ofthe challenges in understanding cellular response to forces is that there is not yet auniversally accepted or fully described instance of a cellular component that respondsspecifically to a particular type of force. Lack of precise molecular mechanisms forforce sensing leaves open the possibility that what is interpreted as a purely mechan-ical stimulus might, in fact, be a chemical stimulus that occurs coincident with theforce application. For example, the effects of shear flow on endothelial and epithelialcells has plausibly been attributed either to a mechanosensor at the apical surface ofthese cells or to chemical sensors that detect concentration differences or gradients insolutes that flow past the cell surface. Similarly, cellular response to forces that deformcell shape can plausibly be described as being due to mechanically induced changes inprotein or membrane molecular conformations or, alternatively, to chemical differ-ences owing to reorientation of the spatial organization of cellular components thatwork cooperatively, with changes due to altering their spatial arrangements but nottheir individual structures.

Several different structures have been proposed as sensors of force. These includeextracellular matrix molecules, the cytoskeleton, transmembrane proteins, proteins atthe membrane-phospholipid interface, elements of the nuclear matrix, chromatin, andthe lipid bilayer itself. The magnitudes of elastic moduli that describe both the cell’sglobal deformation in response to forces and the deformation of specific structuresat the cell surface can help discriminate between different modes of mechanosens-ing. Below we consider the role of the extracellular matrix, matrix receptors, and


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intercellular adhesion receptors in mediating mechanical signals and how this mightin turn impact mechanically activated ion channels.

The Extracellular Matrix in Cell Mechanics

In organs and the connective tissues that envelop organs, cells are embedded in soft ormineralized tissues that provide special situations for the transmission and sensationof mechanical forces. Cells in solid tissues, by virtue of their surrounding proteinmatrices, likely sense and respond to applied forces in ways that are very different,for example, from blood cells flowing past endothelium (52). Connective tissue cellsare therefore mechanically adapted to the rheological properties of the matrix andtheir sensitivity and response to mechanical stimuli are significantly impacted by thematrix proteins that surround them.

Connective tissues provide essential physical and biological properties for trans-mission of gravitational and muscular forces by, for example, functionally joiningmuscle to bone. Another crucial facet of the extracellular matrix is its mechanicalstabilization of tissues subjected to force, a general property that enables tissues andorgans to maintain their shape and to prevent mechanically induced cell damage.Connective tissue matrices are comprised of predominantly collagen fibers and alarge variety of other fibrillar and globular proteins, including fibronectins, laminins,glycosaminoglycans, and tenascin. Specialized mineralizing tissues that exhibit highlyordered hydroxyapatite crystals distributed in a soft extracellular matrix can exhibit awide range of stiffness, as seen in cortical bone compared with cartilage, for example.Notably, fibrillar collagen is the most abundant protein in mammals (53) and in softconnective tissues, it can transmit tensile forces (54) to fibroblasts and many othercell types with appropriate collagen receptors. Based on the structure and composi-tion of tissue-specific proteins, extracellular matrices are highly adapted to transmit,and to protect cells against, different types of mechanical loads, including tensile,compressive, shearing forces, and various combinations thereof (55).

Under conditions of increased loading, connective tissues exhibit clear-cut re-sponses to applied forces, including enhanced remodeling of connective tissue ma-trices (56) and increased proliferation of fibroblasts and osteoblasts, which are theprincipal mesenchymal cell type of soft and mineralized connective tissues, respec-tively. These cells also contribute to the synthesis, degradation, and remodeling ofextracellular matrices, and their sensitivity to mechanical forces enables force-inducedremodeling of the extracellular matrix. Conversely, the mechanical properties of theextracellular matrix have an important influence on the morphology and function ofosteoblasts and fibroblasts (57). For example, mechanically stressed collagen latticeslead to a “synthetic” fibroblast phenotype characterized by induction of connectivetissue synthesis while simultaneously inhibiting matrix degradation (58). Experimentsusing three-dimensional (3-D) collagen gels have also indicated that mechanical load-ing of tissues influences matrix remodeling (59). Thus the extracellular matrix isa conductor of mechanical forces but its structure and composition is also influ-enced by the cellular responses to those same applied or endogenously generatedforces.


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AJ: adherens junction

Mechanotransduction:conversion of a force to achemical or ionic change

Attachment Receptors in Force Transmission and Sensing

Cells in connective tissues adhere to extracellular matrices by a wide variety of matrixreceptors, which may, under certain circumstances, become clustered into the aggre-gates termed focal adhesions or focal complexes. These complexes are candidate sitesfor transfer of contractile forces to the cytoskeleton in both cultured cells and cellsin tissues (60). Because of the facility of growing fibroblasts in vitro and for applyingvarious types of exogenous forces to these cells, cultured fibroblasts have becomeimportant models for exploring mechanosensing and force response mechanisms insolid tissues. In fibroblasts, force transmission is critically dependent on the attach-ment of cells to matrix molecules such as collagen or fibronectin (61); this is manifestalso in force-induced gene expression where tensile forces applied through matrixprotein (but not poly-l-lysine-coated beads) can promote increased expression of theactin-binding protein filamin A (62).

Attachment of connective tissue cells to the extracellular matrix is contingenton the formation and remodeling of integrin-mediated adhesions, but cells also ad-here to each other by intercellular adhesive molecules, such as cadherins, that mayalso act as force sensors (63). As N-cadherin-mediated adherens junctions (AJs) areinfluenced by integrin biology, fibroblasts may be able to integrate mechanical sig-nals from both adhesion systems to coordinate responses relevant to differentiation,organogenesis, and wound healing (64). Several studies have documented mechanicalsignaling responses through both cadherin and integrin adhesions (63, 65). Conse-quently, mechanotransduction, the process by which cells convert mechanical stimuliinto biochemical responses, may not be a finite single process but may instead bea series of interrelated processes that involve the recruitment of a wide variety ofcell attachment, cytoskeletal, and signaling proteins. These proteins may then formdocking and signaling complexes that are oriented in time and space to optimize trans-mission and processing of mechanical signals. Transmembrane proteins, particularlyintegrins and cadherins, are thought to play an important role in the function ofprotein complex–mediated mechanotransduction. Integrins, major transmembraneproteins that connect the cytoskeleton to the extracellular matrix, may be able totransmit mechanical forces across the plasma membrane (66). Because integrins canalso regulate a wide variety of signaling pathways (67), they are well positioned totransduce physical forces into chemical signals.

Transmembrane proteins can activate intracellular biochemical signaling path-ways either by binding an extracellular ligand (chemical signaling) or when they areunfolded or otherwise deformed by force (mechanical signaling). Thus, the adhesivefunction of attachment molecules such as integrins and cadherins is a key elementin mechanosensing and mechanotransduction. When external forces are applied tocultured cells, the cells exert tractional forces on the substratum to which they areattached, for example, focal adhesions complexes (68). The initial reorganization andmaturation of focal complexes into focal adhesions enhance force transmission andalso further increase maturation of the adhesions (69). Conversely, formation of focaladhesions requires tension generated by actin-bound myosin motors within the cell.These cell-generated forces are resisted by substrata if the substrata are sufficiently


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stiff to prevent deformation (70). The processes of force transmission, sensing, andresponse create regulatory opportunities for signaling pathways that may guide cellmigration, matrix assembly, and tissue organization (71) and these opportunities relyheavily on the adhesive domains of cells. The extensive heterogeneity of the mechani-cal properties of cells makes it unlikely that one single general model would be able toexplain cell mechanics (72), particularly in view of the segregated nature of surface ad-hesion receptors on cells (e.g., focal adhesions). Some cases of site-specific responsesthat are heavily influenced by receptor-matrix interactions are difficult to reconcilewith models that posit homogeneous biochemical and mechanical properties. Thispoint is underlined by data showing that application of mechanical forces can be verylocalized and that cellular responses are often restricted to the local application site(73) or in some cases transmitted to discrete sites farther away (74).

Cadherins and Mechanobiology

In mechanically loaded, rapidly remodeling fibrous connective tissues such as theperiodontal ligament, fibroblasts exhibit an extensive network of adherens junctions(AJs) that provide direct physical coupling between adjacent cells (75–78) and endowcells with the capacity to act as a syncitium (63). This property of cells in connectivetissues has received relatively little attention but may be of considerable importancein modeling studies of cell and tissue mechanics. Notably, cadherin-mediated AJsin connective tissue fibroblasts are important for physiological turnover of the ex-tracellular matrix, wound contraction, and intercellular mechanotransduction (76,79–81), processes that involve mechanical signaling processes. Although the forma-tion and maturation of N-cadherin-mediated AJs in response to mechanical loadingare still poorly characterized, new data indicate several interesting insights into cellmechanics.

AJs are adhesive plaques located at the junction of contacting cells, and they aremediated by the classical cadherins, adapter proteins, and actin filaments. AJ-likestructures in solid tissues, such as connective tissues, can be formed by N-cadherin(82, 83), which is broadly expressed by cells in nervous, fibrous, mineralized, muscu-lar, vascular, and adipose tissues. N-cadherin-mediated AJs are of central importancein mechanical aspects of connective tissue function, including the regulation of cellattachment and migration (84–86); wound edge retraction (87); embryonic develop-ment and folding of cell layers (88, 89); and differentiation and formation of neural(90–93), muscle (94–97), bone (98–100), cartilage (101–103), vasculature (104), andfibrous connective tissues (80, 81). All of these processes involve cell mechanics, notonly in interactions with the extracellular matrix but also, importantly, in maintenanceand remodeling of intercellular junctions.

As indicated above, simple, one-dimensional models of mechanical signaling inmany types of tissues do not take into account the remarkable complexity of the to-tal toolbox of components in the extracellular matrix and in intercellular adhesivesystems. In the context of adhesion receptors, cooperative signaling between cad-herin and integrin adhesion systems may be relevant to mechanical signaling and thecoordinated migration of cells that occurs in wound healing. For example, ectopic


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expression of α5 and β1 integrins in primary myoblasts increases N-cadherin ex-pression and contact inhibition (86), implicating integrin signaling in N-cadherin-mediated contact inhibition. In HeLa cells, the importance of paxillin and FAK signal-ing in promoting N-cadherin-mediated contact formation, maintenance, and contactinhibition of motility has been suggested to involve a mechanism employing down-regulation of Rac1 activity at sites subjacent to intercellular contacts (105). Evidently,integrin- and cadherin-mediated adhesions may cooperatively or antagonistically im-pact the formation and maintenance of the opposing adhesion system to regulate cellmechanics. Thus the adhesion of connective tissue cells to various types of extra-cellular matrix proteins can influence the formation of intercellular adhesions and,as a result, the mechanical properties of tissues. For example, epithelial cells adher-ing to fibronectin or laminin maintain intercellular adhesions, whereas, if these cellsattach to collagen, the intercellular junctions dissipate and the cells disperse (106),indicating that different adhesion systems are tightly interdependent in terms of theirmechanical properties. The regulation of cadherin-dependent intercellular junctionsby specific integrin-mediated matrix adhesions has been noted in several differenttypes of cells (107–111), strongly indicating that models predicting mechanical be-haviors of tissues must take into account the cross-talk between the adhesion systemsthat adhere cells to matrix and cells to each other.

Adhesion Systems and the Cytoskeleton in Regulationof Ion Channels

At the single-cell level, mechanical signaling underlies a very large coterie of cellbehaviors, including cell volume control, motility, response to applied forces, andphagocytosis, all of which can produce localized distortions of the cell membrane(112, 113). Force-induced changes of the physical properties of cell membranes maybe able to regulate ion channel conductance (114), directly implicating adhesionsystems to cells and matrix proteins in early events of mechanotransduction. Someexperimental evidence shows that alterations of membrane tension can enhance mem-brane (and presumably ion channel) permeability to calcium; these processes are inturn regulated by the adhesion systems utilized by cells and by the interaction ofadhesion systems with the cytoskeleton. At least one model of mechanotransductionsuggests a central role for the cytoskeleton in regulating membrane ion channels.This hypothesis proposes that cells regulate ion channel conductance, in part, byforce transmission through adhesions to the cytoskeleton, and subsequent cytoskele-tal modification of the gating properties of the channels. As mechanosensitive chan-nels include both stretch-activated and stretch-inactivated channels (114), there isconsiderable scope for how applied forces, adhesion systems, and cytoskeletal pro-teins could impact global membrane permeability. For example, an early event afterN-cadherin ligation of adjacent cells is the activation of plasma membrane stretch-sensitive Ca2+ permeable channels and transient increases of Ca2+ subjacent to sitesof intercellular contact (63). In many types of cells, including connective tissue fi-broblasts, these transients are important determinants of cell contractility, motility(115, 116), and the formation of intercellular adhesions (63, 117). After N-cadherin


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ER: endoplasmic reticulum

ligation, the amplitude of juxtamembrane Ca2+ fluxes correlates temporally with therecruitment of actin assembly at regions of cadherin-mediated adhesions (63, 117).Thus, the mechanics of adhesion formation and remodeling directly impacts fun-damental aspects of cell signaling that impact cell metabolism and are the subjectof numerous studies of mechanical signaling events. The central notion underlyingadhesion system regulation of ion channel conductance is that cell membranes aremajor targets of external mechanical forces (118) and that forces applied to the cellmembrane can be translated into membrane tension. This elevated tension in turnactivates increased ion channel conductance, ion influx, and, potentially, downstreamregulation of gene expression.

As ion transport in various tissues can apparently be regulated by adhesion sys-tems that interact with the cortical actin cytoskeleton, it has been suggested thatstretch-sensitive channels may be able to sense mechanical signals delivered fromthe submembrane, cytoplasmic face of the cell, possibly involving physical linkagesto the cytoskeleton (113). Support for this notion comes from earlier work showingthat transmembrane calcium ion flux is a rapid response to mammalian whole-cellmechanical stretching and is dependent on the relative levels of actin filament as-sembly in individual cells (119). More specifically, using inside-out patch clamping,the involvement of actin assembly in the regulation of nonvoltage-gated sodiumchannels has been demonstrated in human myeloid leukemia cells (120). Stretch-activated channels comprise several functionally characterized ion channels that arepermeable to anions and cations (114, 116), but the molecular definition of thesechannels in mammalian cells has not been completely defined. MscL, a mechanosen-sitive channel in Escherichia coli, was the first major channel molecule to be char-acterized at the molecular level. Thus, MscL may be able to directly contribute tothe “sense” of membrane stretch (121). In mammalian cells, l-type channels may bethe mechanosensor that regulates bone formation (122). Transient receptor potentialchannel proteins have also been cited as possible mechanosensors (123); indeed, workin frog oocytes strongly indicates that TRCPC1 is a mechanosensitive cation channel(124).

Force Transmission to Intracellular Organelles

Although there is a large body of work implicating force transmission throughmembrane-bound adhesion receptors that regulate stretch-sensitive channels in thecell membrane, mechanical stimuli also induce integrin-dependent Ca2+ entry and/orCa2+ release from the endoplasmic reticulum in many cell types, including osteoblasts(125), osteocytes (126), chondrocytes (127, 128), mast cells (129), cardiac myocytes(130), endothelial cells (131), and vascular smooth muscle cells (132, 133). Theseremarkable findings suggest that adhesion receptors, possibly by virtue of their con-nectivity with cytoskeletal elements, can activate calcium release from intracellularstores. One mechanism posits that mechanical stimuli promote release of ATP thatin turn acts on purinergic receptors to discharge Ca2+ from endoplasmic reticu-lum (ER) stores and induce Ca2+ wave propagation (134, 135). However, more re-cent evidence indicates that force activates l-type channels to promote Ca2+ entry


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that stimulates vesicular ATP release (136). Other data from epithelial cells and fi-broblasts have shown that mechanically induced ATP release requires elevations ofintracellular Ca2+ and that Ca2+-dependent exocytosis plays a major role inmechanosensitive ATP release (137). Application of forces to certain cell types pro-motes intracellular release that may rely in part on calcium-induced calcium releasethrough activation of plasma membrane channels (138), activation of a mechanosen-sitive phospholipase A2, or a direct physical connection between the ER and plasmamembrane adhesion receptors (139). Collectively, these data indicate that mechan-ically induced increases of Ca2+ are widely distributed among many different typesof cells, but that the relative importance of mechanically sensitive plasma mem-brane and ER channels is not defined. Recent data indicate that there may belocalized mechanical connections between matrix adhesions on the plasma mem-branes of cells and the endoplasmic reticulum (140, 141), thereby suggesting thepossibility that transmission of mechanical stimuli into the interior of cells can bemediated by physical approximation of intracellular organelles and cell membranemicrodomains.

HOW ARE FORCES EXERTED AT THE CELL SURFACETRANSMITTED THROUGH THE CELL?

The mechanical properties of the cell determine how far and in which direction aforce applied at the cell surface will deform or apply stress to an element withinthe cell. The distribution of forces through a complex viscoelastic material like acell depends on details of the linkage between the site where the force is appliedand the underlying cytoskeleton, the linkage of the membrane to the cytoskeleton,and the continuity of elastic links through the cell interior or linking the membraneto the nucleus. Even prior to active remodeling of the cell in response to the force,details of the preexisting cellular elements determine where in the cell the forcewill be transmitted. For example, when a point force is applied at the membrane to,for example, an integrin that is not tightly coupled to the underlying cortical actinnetwork, the resulting deformation will be largely in the plasma membrane in whichthe integrin is embedded or in elements directly linked to the transmembrane protein.In this case, mechanically sensitive ion channels can be activated, but structures inthe cell interior will not be strained. Any subsequent rearrangement of the internalstructure would occur only secondary to a chemical signal being transduced fromthe mechanical signal. On the other hand, if an integrin is already linked to the 3-Dcytoskeleton, then force will be transmitted directly into the cell interior to deformthe cytoskeleton or move internal organelles. In this case the plasma membrane of thecell might not be stretched, and therefore no influx of Ca2+ would be triggered (142,143). The importance of mechanical reinforcement of integrins for the manner inwhich they signal in response to force underlies the mechanism of mechanoprotection(7, 144), in which repeated or prolonged stress to a transmembrane protein does nottrigger the same response as the initial force application, even though the signalingapparatus remains intact. Similarly, the nature of the contact at the base of the cellaffects how it responds to forces applied elsewhere.


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1.010.840.670.510.340.170.00

X-displacement (µm)

0.670.470.280.08–0.12–0.31–0.51

Z-displacement (µm)

F

F

a

b

x

z

x

z

Figure 1Finite element analysis ofthe displacement field in acell with elastic modulus100 Pa and viscosity100 Pa.s after imposition of500 pN force on a4.5-μm-diameter beadtightly attached to the topof the cell with the depth ofpenetration shown. Onlyone-fourth of the bead isdepicted as the grayquadrant. Thedisplacements predicted 2 safter imposition of stress inthe horizontal (x) andvertical (z) directions areshown in (a) and (b) panels,respectively. Adapted withpermission from Reference145.

Theoretical Predictions of the Stress Field in a Model Cell

Finite element modeling has recently provided maps of how forces applied to thesurface of a cell are transmitted through its interior in the context of magnetic twistingrheometry, a commonly used method (see Table 1) to measure cell surface mechanics.As Figures 1 and 2 show, both the stress and strain fields are transmitted only a fewmicrons away from the point of force application in both viscoelastic and purelyelastic models of cell rheology. Figure 1 shows the displacements in the vertical andhorizontal directions for a model in which approximately one-half of the radius of a4.5 micron sphere is embedded into the membrane of a cell, which is characterized asa viscoelastic body with a shear modulus of 100 Pa and a viscosity of 100 Pa.s (145).The horizontal movements that represent both recoverable elastic displacement andviscous dissipation are significant only a few bead diameters away from the site of the


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x

z

y

a Stress = effective value

b Total deformation = effective value

0.5 µm

x

z

y

Figure 2Finite element analysis ofthe stress (a) and effectivestrain (b) for a 5-μm-highsection of a cell with a4.5 micron beadembedded 10% of itsheight into the cell surfaceand subjected to a 15◦rotation. The cell ismodeled as an elastic bodywith shear modulus1000 Pa and Poisson’sratio = 0.49. Thecolor-coding is maximalfor red (stress = 940 Pa;strain = 0.44) decliningto baseline values at lightblue. Adapted withpermission fromReference 179.

Affine: the macroscopicstrain of a sample is equal tothe microscopic strain atevery point

embedded bead, and extend only a few micrometers into the cell interior. In a purelyelastic cell model with elastic modulus of 1000 Pa (Figure 2), the displacement andcalculated stress fields appear to be even more confined to the region of the embeddedbead. The prediction from these simulations is that cytoskeletal deformation andmovements of embedded organelles should follow a gradient of movements with thelargest magnitudes near the edges of the bead.

Experimental Measurements of Intracellular Strains Causedby Extracellular Forces

Because the interior of the cell, including its cytoskeleton, is anisotropic and ther-modynamically far from equilibrium, the deformation of a cell in response to appliedstress might not correspond to the predicted strain of a homogeneous viscoelasticmaterial. For a simple continuous elastic material, the local strain everywhere withinthe sample corresponds to the macroscopic strain, and such uniform deformationsare called affine. However, many biological materials exhibit non-affine deformation.For example, a detailed study of how the cytoskeleton is deformed by shear stressimaged the vimentin intermediate filament system before and 3 min after impositionof 1.2 Pa shear stress to the dorsal surface of an endothelial cell (146). Reconstructionof network junction points in a triangulation map allowed quantitative measure-ment of the displacement field. As Figure 3 shows, the microscopic displacements ofthe cytoskeleton are highly nonuniform and do not follow closely the direction of thestress. Some junction points even move in a direction opposite to the applied stress.A similar finding that specific sites in the cell interior move significant distances and


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b

aFigure 3(a) Images from an opticalsection of the fluorescentlylabeled vimentin network ofan endothelial cell justbefore (red ) and 3 min after(green) onset of shear stress(1.2 Pa; left to right). Yellowindicates overlapcorresponding to zerofilament displacement.(b) Vertex displacement fieldfrom the inset box in (a).Arrow length andorientation indicate themagnitude and direction ofdisplacement of thecorresponding vertex on theskeleton (gray), shown justbefore flow onset. Adaptedwith permission fromReference 146.

in a direction different from those of forces applied to the cell membrane was madein a study in which twisting motions from a magnetic bead bound to surface integrinslead to displacement of fluorescently labeled mitochondria used as fiducial markersfor the cytoskeleton to which the mitochondria were bound (74, 147). These stud-ies, illustrated in Figure 4, revealed that mitochondria often moved distances and indirections different from what would be expected for application of a local force toa viscoelastic continuum as shown in Figures 1 and 2. Some mitochondria far awayfrom the point of stress moved farther than mitochondria closer to the applied force,and movements toward the force were also noted. Because mitochondria are oftenbound to cytoskeletal filaments, and because the stress carried by individual filamentscan depend strongly on the details of how they are linked to the point of applied


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a b

–20 –10

–10

0

10

20

0

x (µm)

µm Pa

y (µ

m)

10 20

0.06

0.05

0.04

0.03

0.02

0.01

0

250

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150

100

50

0–20 –10

–10

0

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y (µ

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10 20

Figure 4Displacement (a) and stress (b) maps of airway smooth muscle cells after application of 90 Pastress to a 4.5 μm magnetic bead coated with RGD peptides and bound to integrins at the cellsurface. The color map represents the magnitudes of the mitochondria displacements orstresses. The inset in part (b) shows the magnetic bead at the upper right. Adapted withpermission from Reference 74.

force, these measurements are consistent with tensegrity models for cell mechan-ics, which emphasize the geometry of the networks and the interplay of tension andcompression in cytoskeletal structure (148). Some aspects of the differences betweenthe microscopic strains of network nodes and the macroscopic strain are also quali-tatively consistent with recent calculations of the nonaffine strains of dilute networksof cross-linked semiflexible networks, including actin (31, 149) and collagen (150),even when they are not held under tension. Other features, such as the long-rangedisplacement of individual mitochondria after application of a distant force and theeffects of myosin inhibitors, support the concept of tensegrity and implicate the activecontractility of the cell as important in determining how the internally prestressedcytoskeleton responds to external stresses (74, 147).

SENSING AND RESPONDING TO SHEAR STRESS CAUSED BYFLUID FLOW

Characterization of the magnitudes of elastic moduli and the structures at the cellsurface have led to several different mechanisms by which cells may respond to shearstresses caused by fluid flow. As pointed out in a pioneering model of cytoskeletalelasticity based on modeling the filaments as elastic beams, a major challenge tounderstanding how endothelial cells sense shear stress is that the calculated and mea-sured elastic modulus of the cell is on the order of 1000 Pa, but the cell respondsto 0.1 Pa stress at its surface (151). Such a relatively small stress would produce asmall strain the cytoskeleton as a whole. However, the mechanical continuity of thecytoskeleton is required for a decentralized response to the stress and to transmit it


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elsewhere within the cell (152). There is a fairly wide range of values reported forthe Young’s modulus of an endothelial cell, but most studies conclude that the elasticmodulus of the endothelial cytoskeleton is in the range of 100–1000 Pa (Table 1).Therefore, a shear stress of 1 Pa, which is at the high end of the stress reached at thewall of a blood vessel, will strain a cell with an elastic modulus of 1000 Pa only 0.1%.If the typical mesh size of the cytoskeleton is approximately 100 nm, this correspondsto a movement of actin or other filaments of no more than 1 A, a distance almostsurely less than that caused by the random fluctuations from thermal energy. A similarestimate suggests that the stresses caused by fluid flow are also too small to deform astretch-activated ion channel sufficiently to alter its conductivity. Therefore, if the re-sponse to shear flow is really mechanical and not due to flow of chemical messengers,an amplification of the stress due to the fluid flow needs to be hypothesized.

Various structural specializations of flow-sensing cell membranes have been pro-posed as the shear stress sensors. Most models are based on the observation that thesurfaces of cells such as endothelial cells are not smooth (152), and therefore the localstress produced by fluid flow can be significantly higher than the wall shear stresscalculated by classical fluid dynamics (153, 154) for fluid flow across a flat surface.Force imposed at specialized sites can be transmitted through the cytoskeleton tosites in the cell interior (155).

Because of the small forces due to shear flow to which cells respond, the force sen-sor must either be much softer than a typical protein complex or else highly elongatedand projected into the flow field to amplify the force. Two candidate structures toproduce this force amplification are the glycocalyx and the primary cilium. The glyco-calyx, an extended web of large, semiflexible, glycosylated macromolecules extendinglarge distances up from the apical surface of the cell, can amplify the deformationsdue to fluid flow and focus them on the transmembrane proteins to which they areanchored and thereby transduce the shear stress into intracellular signals (156). Thismodality of mechanotransduction is discussed in detail in the chapter by Tarbell et al.in this volume.

Primary Cilium

A convergence of structural and genetic data have recently focused attention on theprimary cilium as a structure responsible for shear stress sensing in some cell types, no-tably epithelial cells and potentially in some endothelial cells (reviewed in References157–161). Motile cilia of, for example, tracheal epithelia generate motion by acti-vation of microtubule motors within an elongated structure formed by a bundle ofnine doublet microtubules surrounding a central pair of microtubules. In contrast,primary cilia are nonmotile because they lack the central microtubules, localize fewermicrotubule motors, and use those motors for intraflagellar transport but not to slidemicrotubules past each other to generate bending movements. Most vertebrate cellsthat face a fluid extracellular compartment, including many endothelial and epithelialcells, contain a single primary cilium that emanates from the mother centriole and ex-tends through the cortical actin meshwork to form membrane-bounded protrusionsthat project as much as 30 microns up from the cell surface. The nonmotile primary


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cilium has been known for 100 years, and has been proposed to be a mechanosensor,largely on the basis of its morphology (reviewed in Reference 162). Molecular ev-idence for the mechanism by which this structure responds to forces has recentlyemerged largely from studies of the dysfunction of renal epithelial cells in polycystickidney disease, and similar or perhaps different molecular strategies may be employedby other cell types that possess primary cilia. Mechanosensing by deflection of themicrotubule-based primary cilium by fluid flow may occur by mechanisms analogousto those by which the actin-based stereocilia of the hair cell detects sound (163).Experimental evidence for mechanical sensing by primary cilia has been reported forrenal and other epithelial cells, osteocytes, and other cell types (see the reviews citedabove), but somewhat surprisingly there is less evidence for such a role in endothelialcells. Several types of endothelial cells express primary cilia (164–168), and flow-induced changes in their structure have been reported in corneal (164) and umbilicalvein endothelia (169); however, in the latter case, physiological levels of fluid shearstress led to their disassembly. On the other hand, they are detected on embryonicendocardial cells in regions of low shear stress (170) on endothelial cells at regionsof atherosclerotic plaques (171), and on hepatic stellate cells (172), all of which areexposed to low levels of shear flow or to disturbed flow.

Bending of the Primary Cilium by Fluid Flow

The deflection of a primary cilium such as that from kangaroo rat kidney epithelialcells shown in Figure 5 subjected to a uniform load perpendicular to the long axis ofthe vertical cilium is given by the equation

d2ψ

ds 2+ k2(ψ) = 0, (1)

where ψ is the tangent angle at a distance s along the cilium contour and k is a constantthat depends on the stiffness of the cilium and the magnitude of the load (i.e., shearstress due to fluid flow). Equation 1 is solvable only by approximation and requiresassumptions about the displacement of the cilium base. Assuming that the ciliumis a uniform nonextensible cylindrical beam and that its base remains fixed at thesurface of the cell, the angle α to which the tip of the cilium bends under a load per unit

a

5 µm

b c d

Figure 5Time sequence of the bending of a primary cilium in response to fluid shear stress. Prior toflow (a), the cilium is vertical and is bent after imposition of low (b) and high (c) levels of shearflow. The cilium regains its resting conformation after flow is stopped (d ). Adapted withpermission from Reference 174.


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length w is given by

a = wL3

6E I, (2)

where L is the length of the cilium, E is its Young’s modulus, and I is the moment ofinertia of the cylindrical cross section. The load per unit length w is related to thefluid flow as (173)

w = 4πρv2d/Re[2.002 − ln(Re)],

where ρ is the density of the fluid, v is the velocity of the fluid, d is the diameter of thecilium and Re is the Reynolds number, estimated to be very low (<2 × 10−5) underthese conditions (174). The diameter of the cilium depends on the cell type and theexperimental conditions and is in the range of 0.12–0.44 micron (166).

The product EI defines the flexural rigidity of the cilium, which has been experi-mentally determined to be 3.1 +/−1 0.8 × 10−23 Nm2 (174) for a bundle containingnine doublet microtubules. The rigidity of the primary cilium derived from imageanalysis in a flow field is surprisingly not much larger than the rigidity of a singlemicrotubule, which is approximately 2.2 × 10−23 Nm2 (175). If these values are bothcorrect, then either doublet microtubules are more flexible than single microtubulesor the primary cilium may have an active mechanism, perhaps involving microtubulemotors, to allow greater flexibility in the microtubule bundle than would be expectedfor the simultaneous bending of nine doublet microtubules.

In principle, there are at least three mechanisms by which the primary cilium orany other long narrow protrusion can amplify forces to allow a cell to respond to verysmall stresses (Figure 6). One mechanism uses the long lever arm of a rigid protrusionto increase the force at the base of the cilium. Extending the cilium several micronsabove the cell surface below a fluid velocity gradient places the tip in a region of theflow field with larger velocity, and therefore more load is put on the protrusion. If thecilium undergoes rigid body rotation, as suggested by some studies (176), the forceat the base of the cilium embedded within the cortical actin meshwork and associatedprotein complexes in the spindle pole body and other microtubule-bound proteinsmay experience large enough forces to produce strains, leading to polypeptide un-folding, release of domain-domain interactions, or other transformations that requirestresses greater than those generated simply by the fluid flowing over a smooth surface.

A second mechanism is suggested by the finding that mechanosensitive ion chan-nels are found away from the base of the cilium at regions where the cilium bends.When a beam of finite diameter bends, the side facing the concave bend is com-pressed and the opposite, outer surface is stretched. Therefore, ion channels and thetransmembrane proteins that regulate them can be subjected to membrane stressessufficient to activate them even if the rest of the cell membrane remains at rest (177).A third mechanism does not require a specific force to be applied to deform the cil-ium, but rather relies on the large elastic extracellular domains of proteins such aspolycystin 1 that might be directly affected by fluid flow. The role of the cilium thenis to project the transmembrane protein away from the cell surface so that it sensesa larger flow field, in a manner analogous to the proposed function of the primary


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Figure 6Schematic diagram of primary cilium and the mechanisms by which it may respond to shearstress. The resting cilium is approximately vertical and is bent by fluid flow above the surfaceof the cell. Embedded in the membrane of the cilium are transmembrane proteins includingion channels, such as polycystin-2, and regulatory proteins, such as polycystin-1, which areassociated with the channel. Bending of the cilium can activate ion channel activity bymechanisms analogous to those by which other stretch-activated channels are regulated.Integrins localized to the distal end of the cilium may similarly be activated by deformationsproduced when the cilium is displaced. Alternatively, the bending of the cilium can amplifyforce transmitted to the base of the cilium embedded in the cell interior because of the rigidityof the microtubule core and deformation of proteins at the base of the cilium may transducethe mechanical signal.

cilium as a chemical sensor that places chemical receptors away from the cell surface.The potential for the primary cilium to act as a force sensor may not be limited to fluidshear stress, as integrins have recently been identified to localize to the primary cilia ofchondrocytes and may therefore act in cell-matrix or cell-cell mechanosensing (178).

CONCLUSIONS

A survey of recent studies shows that many factors, including but not limited to thecytoskeleton, determine the mechanical properties of the cell. The strain-stiffening


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nature of the cytoskeleton allows for stress-dependent cell stiffening and reinforce-ment of cell rigidity at points where mechanical forces are applied. The magnitude ofelastic moduli of a variety of cell types have been measured, and as the experimentalsystems and methods to derive elastic moduli from force-displacement measurementsimprove, the range of elastic moduli reported by different methods appears to be con-verging to the range of 100 to 10,000 Pa for measurements made at modest strainsfor periods of seconds. At the same time, it is also clear that cell stiffness is a het-erogeneous parameter that varies spatially and temporally as cells respond to theirenvironment. The mechanical features of cells allow them to respond to a range ofphysical forces, and quantitative measures of the magnitudes of force required forcell activation and the elastic moduli of the cells that respond will help elucidatemechanisms of mechanotransduction that regulate many aspects of cell function.

SUMMARY POINTS

1. Single cell measurements typically yield values in the range of 100 to10,000 Pa for cell elastic moduli.

2. Cell stiffness increases as cells develop internal tension.

3. Cell-cell contacts are as important as cell-matrix interfaces as sites ofmechanotransduction.

4. Elongated structures, such as the glycocalyx and the primary cilium, mayenable cells to sense the small stresses generated by fluid flow.

5. The time-dependence of cell rheology reveals features that differ from thoseof isotropic biopolymer gels.

DISCLOSURE STATEMENT

The authors are not aware of any biases that might be perceived as affecting theobjectivity of this review.

LITERATURE CITED

1. Roesel von Rosenhof AJ. 1744. Der kleine Proteus. Der Insecten-Belustigung.Nurenberg 3:622–24

2. Heilbrunn L. 1956. The Dynamics of Living Protoplasm. New York: Academic.327 pp.

3. Freundlich H, Seifriz W. 1922. Ueber die elastizitat der Solen und Gelen.Zeitschr. Phys. Chem. 104:233

4. Seifriz W. 1923. An elastic value of protoplasm, with further observations onthe viscosity of protoplasm. J. Exp. Biol. 2:1–11

5. Stossel TP. 1993. On the crawling of animal cells. Science 260:1086–946. Vogel V, Sheetz M. 2006. Local force and geometry sensing regulate cell func-

tions. Nat. Rev. Mol. Cell Biol. 7:265–75


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

7. Bershadsky A, Kozlov M, Geiger B. 2006. Adhesion-mediated mechanosensitiv-ity: a time to experiment, and a time to theorize. Curr. Opin. Cell Biol. 18:472–81

8. Lieber SC, Aubry N, Pain J, Diaz G, Kim SJ, Vatner SF. 2004. Aging increasesstiffness of cardiac myocytes measured by atomic force microscopy nanoinden-tation. Am. J. Physiol. Heart Circ. Physiol. 287:H645–51

9. Levental I, Georges PC, Janmey PA. 2007. Soft biological materials and theirimpact on cell function. Soft Matter 1:299–306

10. Janmey PA, Euteneuer U, Traub P, Schliwa M. 1991. Viscoelastic properties ofvimentin compared with other filamentous biopolymer networks. J. Cell Biol.113:155–60

11. Takai E, Landesberg R, Katz RW, Hung CT, Guo XE. 2006. Substrate mod-ulation of osteoblast adhesion strength, focal adhesion kinase activation, andresponsiveness to mechanical stimuli. Mol. Cell Biomech. 3:1–12

12. Trickey WR, Vail TP, Guilak F. 2004. The role of the cytoskeleton in theviscoelastic properties of human articular chondrocytes. J. Orthop. Res. 22:131–39

13. Costa KD, Sim AJ, Yin FC. 2006. Non-Hertzian approach to analyzing me-chanical properties of endothelial cells probed by atomic force microscopy. J.Biomech. Eng. 128:176–84

14. Wachsstock DH, Schwartz WH, Pollard TD. 1993. Affinity of alpha-actininfor actin determines the structure and mechanical properties of actin filamentgels. Biophys. J. 65:205–14

15. Gardel ML, Nakamura F, Hartwig JH, Crocker JC, Stossel TP, Weitz DA.2006. Prestressed F-actin networks cross-linked by hinged filamins replicatemechanical properties of cells. Proc. Natl. Acad. Sci. USA 103:1762–67

16. Janmey PA, Hvidt S, Lamb J, Stossel TP. 1990. Resemblance of actin-bindingprotein/actin gels to covalently crosslinked networks. Nature 345:89–92

17. MacKintosh FC, Kas J, Janmey PA. 1995. Elasticity of semiflexible biopolymernetworks. Phys. Rev. Lett. 75:4425–28

18. Mast SO. 1923. Mechanics of locomotion in Amoeba. Proc. Natl. Acad. Sci. USA9:258–61

19. Humphrey D, Duggan C, Saha D, Smith D, Kas J. 2002. Active fluidization ofpolymer networks through molecular motors. Nature 416:413–16

20. Guck J, Schinkinger S, Lincoln B, Wottawah F, Ebert S, et al. 2005. Opticaldeformability as an inherent cell marker for testing malignant transformationand metastatic competence. Biophys. J. 88:3689–98

21. Fernandez P, Pullarkat PA, Ott A. 2006. A master relation defines the nonlinearviscoelasticity of single fibroblasts. Biophys. J. 90:3796–805

22. Kaibara M, Fukada E. 1970. Dynamic viscoelastic study for the structure offibrin networks in the clots of blood and plasma. Biorheology 6:329–39

23. Roberts WW, Kramer O, Rosser RW, Nestler FH, Ferry JD. 1974. Rheologyof fibrin clots. I. Dynamic viscoelastic properties and fluid permeation. Biophys.Chem. 1:152–60

24. Ryan EA, Mockros LF, Weisel JW, Lorand L. 1999. Structural origins of fibrinclot rheology. Biophys. J. 77:2813–26


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

25. Shah JV, Janmey PA. 1997. Strain hardening of fibrin gels and plasma clots.Rheol. Acta 36:262–68

26. Djabourov M, Lechaire JP, Gaill F. 1993. Structure and rheology of gelatin andcollagen gels. Biorheology 30:191–205

27. Morse DC. 2001. Tube diameter in tightly entangled solutions of semiflexiblepolymers. Phys. Rev. E Stat. Nonlin. Soft Matter Phys. 63:031502

28. Storm C, Pastore JJ, MacKintosh FC, Lubensky TC, Janmey PA. 2005. Non-linear elasticity in biological gels. Nature 435:191–94

29. Dorrington KL, McCrum NG. 1977. Elastin as a rubber. Biopolymers 16:1201–22

30. Gosline J, Lillie M, Carrington E, Guerette P, Ortlepp C, Savage K. 2002.Elastic proteins: biological roles and mechanical properties. Philos. Trans. R.Soc. London B Biol. Sci. 357:121–32

31. Onck PR, Koeman T, van Dillen T, van der Giessen E. 2005. Alternative ex-planation of stiffening in cross-linked semiflexible networks. Phys. Rev. Lett.95:178102

32. dos Remedios CG, Chhabra D, Kekic M, Dedova IV, Tsubakihara M, et al.2003. Actin binding proteins: regulation of cytoskeletal microfilaments. Physiol.Rev. 83:433–73

33. Winder SJ, Ayscough KR. 2005. Actin-binding proteins. J. Cell Sci. 118:651–5434. Keller M, Tharmann R, Dichtl MA, Bausch AR, Sackmann E. 2003. Slow fila-

ment dynamics and viscoelasticity in entangled and active actin networks. Philos.Trans. A Math. Phys. Eng. Sci. 361:699–711; discuss. 712

35. Kruse K, Joanny JF, Julicher F, Prost J, Sekimoto K. 2005. Generic theoryof active polar gels: a paradigm for cytoskeletal dynamics. Eur. Phys. J. E SoftMatter 16:5–16

36. Voituriez R, Joanny JF, Prost J. 2006. Generic phase diagram of active polarfilms. Phys. Rev. Lett. 96:028102

37. Pourati J, Maniotis A, Spiegel D, Schaffer JL, Butler JP, et al. 1998. Is cytoskele-tal tension a major determinant of cell deformability in adherent endothelialcells? Am. J. Physiol. Cell Physiol. 274:C1283–89

38. Smith BA, Tolloczko B, Martin JG, Grutter P. 2005. Probing the viscoelasticbehavior of cultured airway smooth muscle cells with atomic force microscopy:stiffening induced by contractile agonist. Biophys. J. 88:2994–3007

39. Trepat X, Grabulosa M, Buscemi L, Rico F, Farre R, Navajas D. 2005. Throm-bin and histamine induce stiffening of alveolar epithelial cells. J. Appl. Physiol.98:1567–74

40. Trepat X, Grabulosa M, Puig F, Maksym GN, Navajas D, Farre R. 2004. Vis-coelasticity of human alveolar epithelial cells subjected to stretch. Am. J. Physiol.Lung Cell Mol. Physiol. 287:L1025–34

41. Wang N, Stamenovic D. 2000. Contribution of intermediate filaments to cellstiffness, stiffening, and growth. Am. J. Physiol. Cell Physiol. 279:C188–94

42. Lau AW, Hoffman BD, Davies A, Crocker JC, Lubensky TC. 2003. Microrhe-ology, stress fluctuations, and active behavior of living cells. Phys. Rev. Lett.91:198101


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

43. Heilbrunn LV. 1924. The colloid chemistry of protoplasm: IV. The heat coag-ulation of protoplasm. Am. J. Physiol. 69:190–99

44. Hoffman BD, Massiera G, Van Citters KM, Crocker JC. 2006. The consensusmechanics of cultured mammalian cells. Proc. Natl. Acad. Sci. USA 103:10259–64

45. Lim CT, Zhou EH, Quek ST. 2006. Mechanical models for living cells—areview. J. Biomech. 39:195–216

46. Alcaraz J, Buscemi L, Grabulosa M, Trepat X, Fabry B, et al. 2003. Microrheol-ogy of human lung epithelial cells measured by atomic force microscopy. Biophys.J. 84:2071–79

47. Fabry B, Maksym GN, Butler JP, Glogauer M, Navajas D, et al. 2003. Timescale and other invariants of integrative mechanical behavior in living cells. Phys.Rev. E Stat. Nonlin. Soft Matter Phys. 68:041914

48. Puig-de-Morales M, Millet E, Fabry B, Navajas D, Wang N, et al. 2004. Cy-toskeletal mechanics in adherent human airway smooth muscle cells: probespecificity and scaling of protein-protein dynamics. Am. J. Physiol. Cell Physiol.287:C643–54

49. Stamenovic D, Suki B, Fabry B, Wang N, Fredberg JJ. 2004. Rheology ofairway smooth muscle cells is associated with cytoskeletal contractile stress. J.Appl. Physiol. 96:1600–5

50. Laudadio RE, Millet EJ, Fabry B, An SS, Butler JP, Fredberg JJ. 2005. Rat airwaysmooth muscle cell during actin modulation: rheology and glassy dynamics. Am.J. Physiol. Cell Physiol. 289:C1388–95

51. Deng L, Trepat X, Butler JP, Millet E, Morgan KG, et al. 2006. Fast and slowdynamics of the cytoskeleton. Nat. Mater. 5:636–40

52. Tzima E, Irani-Tehrani M, Kiosses WB, Dejana E, Schultz DA, et al. 2005. Amechanosensory complex that mediates the endothelial cell response to fluidshear stress. Nature 437:426–31

53. Perez-Tamayo R. 1978. Pathology of collagen degradation. A review. Am. J.Pathol. 92:508–66

54. Provenzano PP, Vanderby RJ. 2006. Collagen fibril morphology and organiza-tion: implications for force transmission in ligament and tendon. Matrix Biol.25:71–84

55. Warden SJ, Hurst JA, Sanders MS, Turner CH, Burr DB, Li J. 2005. Bone adap-tation to a mechanical loading program significantly increases skeletal fatigueresistance. J. Bone Miner. Res. 20:809–16

56. Ozaki S, Kaneko S, Podyma-Inoue KA, Yanagishita M, Soma K. 2005. Mod-ulation of extracellular matrix synthesis and alkaline phosphatase activity ofperiodontal ligament cells by mechanical stress. J. Periodontal. Res. 40:110–17

57. Hinz B, Gabbiani G. 2003. Mechanisms of force generation and transmissionby myofibroblasts. Curr. Opin. Biotechnol. 14:538–46

58. Kessler D, Dethlefsen S, Haase I, Plomann M, Hirche F, et al. 2001. Fibroblastsin mechanically stressed collagen lattices assume a “synthetic” phenotype. J. Biol.Chem. 276:36575–85

59. Mudera VC, Pleass R, Eastwood M, Tarnuzzer R, Schultz G, et al. 2000. Molec-ular responses of human dermal fibroblasts to dual cues: contact guidance andmechanical load. Cell Motil. Cytoskel. 45:1–9


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

60. Ingber DE. 2003. Tensegrity II. How structural networks influence cellularinformation processing networks. J. Cell Sci. 116:1397–408

61. Hinz B. 2006. Masters and servants of the force: the role of matrix adhesions inmyofibroblast force perception and transmission. Eur. J. Cell Biol. 85:175–81

62. D’Addario M, Arora PD, Ellen RP, McCulloch CA. 2002. Interaction of p38and Sp1 in a mechanical force-induced, beta 1 integrin-mediated transcrip-tional circuit that regulates the actin-binding protein filamin-A. J. Biol. Chem.277:47541–50

63. Ko KS, Arora PD, McCulloch CA. 2001. Cadherins mediate intercellularmechanical signaling in fibroblasts by activation of stretch-sensitive calcium-permeable channels. J. Biol. Chem. 276:35967–77

64. Linask KK, Manisastry S, Han M. 2005. Cross talk between cell-cell and cell-matrix adhesion signaling pathways during heart organogenesis: implicationsfor cardiac birth defects. Microsc. Microanal. 11:200–8

65. Potard US, Butler JP, Wang N. 1997. Cytoskeletal mechanics in confluentepithelial cells probed through integrins and E-cadherins. Am. J. Physiol. CellPhysiol. 272:C1654–63

66. Huang S, Ingber DE. 1999. The structural and mechanical complexity of cell-growth control. Nat. Cell Biol. 1:E131–38

67. Schwartz MA, Assoian RK. 2001. Integrins and cell proliferation: regulationof cyclin-dependent kinases via cytoplasmic signaling pathways. J. Cell Sci.114:2553–60

68. Schoenwaelder SM, Burridge K. 1999. Bidirectional signaling between the cy-toskeleton and integrins. Curr. Opin. Cell Biol. 11:274–86

69. Galbraith CG, Yamada KM, Sheetz MP. 2002. The relationship between forceand focal complex development. J. Cell Biol. 159:695–705

70. Pelham RJJ, Wang Y. 1997. Cell locomotion and focal adhesions are regulatedby substrate flexibility. Proc. Natl. Acad. Sci. USA 94:13661–65

71. Katsumi A, Orr AW, Tzima E, Schwartz MA. 2004. Integrins in mechanotrans-duction. J. Biol. Chem. 279:12001–4

72. Heidemann SR, Wirtz D. 2004. Towards a regional approach to cell mechanics.Trends Cell Biol. 14:160–66

73. Bershadsky AD, Balaban NQ, Geiger B. 2003. Adhesion-dependent cellmechanosensitivity. Annu. Rev. Cell Dev. Biol. 19:677–95

74. Hu S, Chen J, Fabry B, Numaguchi Y, Gouldstone A, et al. 2003. Intracellularstress tomography reveals stress focusing and structural anisotropy in cytoskele-ton of living cells. Am. J. Physiol. Cell Physiol. 285:C1082–90

75. Beertsen W, Everts V. 1980. Junctions between fibroblasts in mouse periodontalligament. J. Periodontal. Res. 15:655–68

76. Ko KS, McCulloch CA. 2001. Intercellular mechanotransduction: cellular cir-cuits that coordinate tissue responses to mechanical loading. Biochem. Biophys.Res. Commun. 285:1077–83

77. Luo Y, Radice GL. 2003. Cadherin-mediated adhesion is essential for myofibrilcontinuity across the plasma membrane but not for assembly of the contractileapparatus. J. Cell Sci. 116:1471–79


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

78. Shore RC, Berkovitz BK, Moxham BJ. 1981. Intercellular contacts betweenfibroblasts in the periodontal connective tissues of the rat. J. Anat. 133:67–76

79. Hinz B, Gabbiani G. 2003. Cell-matrix and cell-cell contacts of myofibroblasts:role in connective tissue remodeling. Thromb. Haemost. 90:993–1002

80. Hinz B, Pittet P, Smith-Clerc J, Chaponnier C, Meister JJ. 2004. Myofibroblastdevelopment is characterized by specific cell-cell adherens junctions. Mol. Biol.Cell 15:4310–20

81. Tomasek JJ, Gabbiani G, Hinz B, Chaponnier C, Brown RA. 2002. Myofibrob-lasts and mechano-regulation of connective tissue remodelling. Nat. Rev. Mol.Cell Biol. 3:349–63

82. Ko K, Arora P, Lee W, McCulloch C. 2000. Biochemical and functional char-acterization of intercellular adhesion and gap junctions in fibroblasts. Am. J.Physiol. Cell Physiol. 279:C147–57

83. Heaysman JE, Pegrum SM. 1973. Early contacts between fibroblasts. An ultra-structural study. Exp. Cell Res. 78:71–78

84. Akitaya T, Bronner-Fraser M. 1992. Expression of cell adhesion moleculesduring initiation and cessation of neural crest cell migration. Dev. Dyn. 194:12–20

85. Bronner-Fraser M, Wolf JJ, Murray BA. 1992. Effects of antibodies against N-cadherin and N-CAM on the cranial neural crest and neural tube. Dev. Biol.153:291–301

86. Huttenlocher A, Lakonishok M, Kinder M, Wu S, Truong T, et al. 1998. Inte-grin and cadherin synergy regulates contact inhibition of migration and motileactivity. J. Cell Biol. 141:515–26

87. De Wever O, Westbroek W, Verloes A, Bloemen N, Bracke M, et al. 2004.Critical role of N-cadherin in myofibroblast invasion and migration in vitrostimulated by colon-cancer-cell-derived TGF-beta or wounding. J. Cell Sci.117:4691–703

88. Garcia-Castro MI, Vielmetter E, Bronner-Fraser M. 2000. N-cadherin, a celladhesion molecule involved in establishment of embryonic left-right asymme-try. Science 288:1047–51

89. Radice GL, Rayburn H, Matsunami H, Knudsen KA, Takeichi M, Hynes RO.1997. Developmental defects in mouse embryos lacking N-cadherin. Dev. Biol.181:64–78

90. Okamura K, Tanaka H, Yagita Y, Saeki Y, Taguchi A, et al. 2004. Cadherinactivity is required for activity-induced spine remodeling. J. Cell Biol. 167:961–72

91. Yu X, Malenka RC. 2003. Beta-catenin is critical for dendritic morphogenesis.Nat. Neurosci. 6:1169–77

92. Uchida N, Honjo Y, Johnson KR, Wheelock MJ, Takeichi M. 1996. Thecatenin/cadherin adhesion system is localized in synaptic junctions borderingtransmitter release zones. J. Cell Biol. 135:767–79

93. Bamji SX. 2005. Cadherins: actin with the cytoskeleton to form synapses. Neuron47:175–78

94. Goichberg P, Geiger B. 1998. Direct involvement of N-cadherin-mediatedsignaling in muscle differentiation. Mol. Biol. Cell 9:3119–31


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

95. Redfield A, Nieman MT, Knudsen KA. 1997. Cadherins promote skeletal mus-cle differentiation in three-dimensional cultures. J. Cell Biol. 138:1323–31

96. George-Weinstein M, Gerhart J, Blitz J, Simak E, Knudsen KA. 1997. N-cadherin promotes the commitment and differentiation of skeletal muscle pre-cursor cells. Dev. Biol. 185:14–24

97. Mege RM, Goudou D, Diaz C, Nicolet M, Garcia L, et al. 1992. N-cadherinand N-CAM in myoblast fusion: compared localisation and effect of blockadeby peptides and antibodies. J. Cell Sci. 103(Pt. 4):897–906

98. Castro CH, Shin CS, Stains JP, Cheng SL, Sheikh S, et al. 2004. Targetedexpression of a dominant-negative N-cadherin in vivo delays peak bone massand increases adipogenesis. J. Cell Sci. 117:2853–64

99. Cheng SL, Shin CS, Towler DA, Civitelli R. 2000. A dominant negative cadherininhibits osteoblast differentiation. J. Bone Miner. Res. 15:2362–70

100. Lemonnier J, Hay E, Delannoy P, Lomri A, Modrowski D, et al. 2001. Role ofN-cadherin and protein kinase C in osteoblast gene activation induced by theS252W fibroblast growth factor receptor 2 mutation in Apert craniosynostosis.J. Bone Miner. Res. 16:832–45

101. Haas AR, Tuan RS. 1999. Chondrogenic differentiation of murine C3H10T1/2multipotential mesenchymal cells: II. Stimulation by bone morphogeneticprotein-2 requires modulation of N-cadherin expression and function. Differ-entiation 64:77–89

102. Oberlender SA, Tuan RS. 1994. Spatiotemporal profile of N-cadherin expres-sion in the developing limb mesenchyme. Cell Adhes. Commun. 2:521–37

103. Tuli R, Tuli S, Nandi S, Huang X, Manner PA, et al. 2003. Transforming growthfactor-beta-mediated chondrogenesis of human mesenchymal progenitor cellsinvolves N-cadherin and mitogen-activated protein kinase and Wnt signalingcross-talk. J. Biol. Chem. 278:41227–36

104. Luo Y, Radice GL. 2005. N-cadherin acts upstream of VE-cadherin in control-ling vascular morphogenesis. J. Cell Biol. 169:29–34

105. Yano H, Mazaki Y, Kurokawa K, Hanks SK, Matsuda M, Sabe H. 2004. Rolesplayed by a subset of integrin signaling molecules in cadherin-based cell-celladhesion. J. Cell Biol. 166:283–95

106. Sander EE, van Delft S, ten Klooster JP, Reid T, van der Kammen RA, et al.1998. Matrix-dependent Tiam1/Rac signaling in epithelial cells promotes eithercell-cell adhesion or cell migration and is regulated by phosphatidylinositol 3-kinase. J. Cell Biol. 143:1385–98

107. Clark P. 1994. Modulation of scatter factor/hepatocyte growth factor activityby cell-substratum adhesion. J. Cell Sci. 107(Pt. 5):1265–75

108. Kawano K, Kantak SS, Murai M, Yao CC, Kramer RH. 2001. Integrin al-pha3beta1 engagement disrupts intercellular adhesion. Exp. Cell Res. 262:180–96

109. Marsden M, DeSimone DW. 2003. Integrin-ECM interactions regulatecadherin-dependent cell adhesion and are required for convergent extensionin Xenopus. Curr. Biol. 13:1182–91

110. Sakai T, Larsen M, Yamada KM. 2003. Fibronectin requirement in branchingmorphogenesis. Nature 423:876–81


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

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ownl

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s.an

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org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

111. Gimond C, van Der Flier A, van Delft S, Brakebusch C, Kuikman I, et al. 1999.Induction of cell scattering by expression of beta1 integrins in beta1-deficientepithelial cells requires activation of members of the rho family of GTPases anddownregulation of cadherin and catenin function. J. Cell Biol. 147:1325–40

112. Tavernarakis N, Driscoll M. 2001. Mechanotransduction in Caenorhabditis ele-gans: the role of DEG/ENaC ion channels. Cell Biochem. Biophys. 35:1–18

113. Alenghat FJ, Ingber DE. 2002. Mechanotransduction: all signals point to cy-toskeleton, matrix, and integrins. Sci. STKE 2002:PE6

114. Morris CE. 1990. Mechanosensitive ion channels. J. Membr. Biol. 113:93–107115. Doyle AD, Lee J. 2005. Cyclic changes in keratocyte speed and traction stress

arise from Ca2+-dependent regulation of cell adhesiveness. J. Cell Sci. 118:369–79

116. Lee J, Ishihara A, Oxford G, Johnson B, Jacobson K. 1999. Regulation of cellmovement is mediated by stretch-activated calcium channels. Nature 400:382–86

117. Ko KS, Arora PD, Bhide V, Chen A, McCulloch CA. 2001. Cell-cell adhesionin human fibroblasts requires calcium signaling. J. Cell Sci. 114:1155–67

118. Martinac B. 2004. Mechanosensitive ion channels: molecules of mechanotrans-duction. J. Cell Sci. 117:2449–60

119. Arora PD, McCulloch CA. 1994. Dependence of collagen remodelling on alpha-smooth muscle actin expression by fibroblasts. J. Cell Physiol. 159:161–75

120. Shumilina EV, Negulyaev YA, Morachevskaya EA, Hinssen H, Khaitlina SY.2003. Regulation of sodium channel activity by capping of actin filaments. Mol.Biol. Cell 14:1709–16

121. Sukharev SI, Blount P, Martinac B, Kung C. 1997. Mechanosensitive channelsof Escherichia coli: the MscL gene, protein, and activities. Annu. Rev. Physiol.59:633–57

122. Li J, Duncan RL, Burr DB, Turner CH. 2002. L-type calcium channels mediatemechanically induced bone formation in vivo. J. Bone Miner. Res. 17:1795–800

123. Minke B, Cook B. 2002. TRP channel proteins and signal transduction. Physiol.Rev. 82:429–72

124. Maroto R, Raso A, Wood TG, Kurosky A, Martinac B, Hamill OP. 2005.TRPC1 forms the stretch-activated cation channel in vertebrate cells. Nat. CellBiol. 7:179–85

125. Haut Donahue TL, Genetos DC, Jacobs CR, Donahue HJ, Yellowley CE.2004. Annexin V disruption impairs mechanically induced calcium signaling inosteoblastic cells. Bone 35:656–63

126. Mikuni-Takagaki Y. 1999. Mechanical responses and signal transduction path-ways in stretched osteocytes. J. Bone Miner. Metab. 17:57–60

127. Lee HS, Millward-Sadler SJ, Wright MO, Nuki G, Salter DM. 2000. Integrinand mechanosensitive ion channel-dependent tyrosine phosphorylation of fo-cal adhesion proteins and beta-catenin in human articular chondrocytes aftermechanical stimulation. J. Bone Miner. Res. 15:1501–9

128. Mobasheri A, Carter SD, Martin-Vasallo P, Shakibaei M. 2002. Integrins andstretch activated ion channels; putative components of functional cell surfacemechanoreceptors in articular chondrocytes. Cell Biol. Int. 26:1–18


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

129. Stokes AJ, Shimoda LM, Koblan-Huberson M, Adra CN, Turner H. 2004.A TRPV2-PKA signaling module for transduction of physical stimuli in mastcells. J. Exp. Med. 200:137–47

130. Lammerding J, Kamm RD, Lee RT. 2004. Mechanotransduction in cardiacmyocytes. Ann. N.Y. Acad. Sci. 1015:53–70

131. Nilius B, Droogmans G, Wondergem R. 2003. Transient receptor potentialchannels in endothelium: solving the calcium entry puzzle? Endothelium 10:5–15

132. Davis MJ, Wu X, Nurkiewicz TR, Kawasaki J, Davis GE, et al. 2001. Integrinsand mechanotransduction of the vascular myogenic response. Am. J. Physiol.Heart Circ. Physiol. 280:H1427–33

133. Wu X, Mogford JE, Platts SH, Davis GE, Meininger GA, Davis MJ. 1998.Modulation of calcium current in arteriolar smooth muscle by alphav beta3 andalpha5 beta1 integrin ligands. J. Cell Biol. 143:241–52

134. Sauer H, Hescheler J, Wartenberg M. 2000. Mechanical strain-induced Ca(2+)waves are propagated via ATP release and purinergic receptor activation. Am.J. Physiol. Cell Physiol. 279:C295–307

135. Romanello M, D’Andrea P. 2001. Dual mechanism of intercellular communi-cation in HOBIT osteoblastic cells: a role for gap-junctional hemichannels. J.Bone Miner. Res. 16:1465–76

136. Genetos DC, Geist DJ, Liu D, Donahue HJ, Duncan RL. 2005. Fluid shear-induced ATP secretion mediates prostaglandin release in MC3T3-E1 os-teoblasts. J. Bone Miner. Res. 20:41–49

137. Boudreault F, Grygorczyk R. 2004. Cell swelling-induced ATP release is tightlydependent on intracellular calcium elevations. J. Physiol. 561:499–513

138. Niggel J, Sigurdson W, Sachs F. 2000. Mechanically induced calcium move-ments in astrocytes, bovine aortic endothelial cells and C6 glioma cells. J. Membr.Biol. 174:121–34

139. Hamill OP, Martinac B. 2001. Molecular basis of mechanotransduction in livingcells. Physiol. Rev. 81:685–740

140. Wang Q, Downey GP, Herrera-Abreu MT, Kapus A, McCulloch CA. 2005.SHP-2 modulates interleukin-1-induced Ca2+ flux and ERK activation viaphosphorylation of phospholipase Cgamma1. J. Biol. Chem. 280:8397–406

141. Wang Q, Herrera Abreu MT, Siminovitch K, Downey GP, McCulloch CA.2006. Phosphorylation of SHP-2 regulates interactions between the endoplas-mic reticulum and focal adhesions to restrict IL-1-induced Ca2+ signaling. J.Biol. Chem. 281:31093–105

142. Glogauer M, Arora P, Chou D, Janmey PA, Downey GP, McCulloch CA. 1998.The role of actin-binding protein 280 in integrin-dependent mechanoprotec-tion. J. Biol. Chem. 273:1689–98

143. Glogauer M, Arora P, Yao G, Sokholov I, Ferrier J, McCulloch CA. 1997.Calcium ions and tyrosine phosphorylation interact coordinately with actin toregulate cytoprotective responses to stretching. J. Cell Sci. 110(Pt. 1):11–21

144. von Wichert G, Haimovich B, Feng GS, Sheetz MP. 2003. Force-dependentintegrin-cytoskeleton linkage formation requires downregulation of focal com-plex dynamics by Shp2. EMBO J. 22:5023–35


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

145. Karcher H, Lammerding J, Huang H, Lee RT, Kamm RD, Kaazempur-MofradMR. 2003. A three-dimensional viscoelastic model for cell deformation withexperimental verification. Biophys. J. 85:3336–49

146. Helmke BP, Rosen AB, Davies PF. 2003. Mapping mechanical strain of anendogenous cytoskeletal network in living endothelial cells. Biophys. J. 84:2691–99

147. Hu S, Wang N. 2006. Control of stress propagation in the cytoplasm by prestressand loading frequency. Mol. Cell Biomech. 3:49–60

148. Ingber DE. 2006. Cellular mechanotransduction: putting all the pieces togetheragain. FASEB J. 20:811–27

149. Head DA, Levine AJ, MacKintosh FC. 2003. Distinct regimes of elastic responseand deformation modes of cross-linked cytoskeletal and semiflexible polymernetworks. Phys. Rev. E Stat. Nonlin. Soft Matter Phys. 68:061907

150. Chandran PL, Barocas VH. 2006. Affine versus nonaffine fibril kinematics incollagen networks: theoretical studies of network behavior. J. Biomech. Eng.128:259–70

151. Satcher RLJ, Dewey CFJ. 1996. Theoretical estimates of mechanical propertiesof the endothelial cell cytoskeleton. Biophys. J. 71:109–18

152. Davies PF, Mundel T, Barbee KA. 1995. A mechanism for heterogeneous en-dothelial responses to flow in vivo and in vitro. J. Biomech. 28:1553–60

153. Davies PF, Spaan JA, Krams R. 2005. Shear stress biology of the endothelium.Ann. Biomed. Eng. 33:1714–18

154. Helmke BP, Davies PF. 2002. The cytoskeleton under external fluid mechan-ical forces: hemodynamic forces acting on the endothelium. Ann. Biomed. Eng.30:284–96

155. Li YS, Haga JH, Chien S. 2005. Molecular basis of the effects of shear stress onvascular endothelial cells. J. Biomech. 38:1949–71

156. Florian JA, Kosky JR, Ainslie K, Pang Z, Dull RO, Tarbell JM. 2003. Heparansulfate proteoglycan is a mechanosensor on endothelial cells. Circ. Res. 93:e136-42

157. Nauli SM, Zhou J. 2004. Polycystins and mechanosensation in renal and nodalcilia. Bioessays 26:844–56

158. Praetorius HA, Spring KR. 2005. A physiological view of the primary cilium.Annu. Rev. Physiol. 67:515–29

159. Singla V, Reiter JF. 2006. The primary cilium as the cell’s antenna: signaling ata sensory organelle. Science 313:629–33

160. Whitfield JF. 2003. Primary cilium—is it an osteocyte’s strain-sensing flowme-ter? J. Cell Biochem. 89:233–37

161. Pazour GJ, Witman GB. 2003. The vertebrate primary cilium is a sensoryorganelle. Curr. Opin. Cell Biol. 15:105–10

162. Wheatley DN. 2005. Landmarks in the first hundred years of primary (9+0)cilium research. Cell Biol. Int. 29:333–39

163. Weinbaum S, Guo P, You L. 2001. A new view of mechanotransduction andstrain amplification in cells with microvilli and cell processes. Biorheology 38:119–42


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

164. Doughty MJ. 1998. Changes in cell surface primary cilia and microvilli concur-rent with measurements of fluid flow across the rabbit corneal endothelium exvivo. Tiss. Cell 30:634–43

165. Huang BQ, Masyuk TV, Muff MA, Tietz PS, Masyuk AI, Larusso NF. 2006.Isolation and characterization of cholangiocyte primary cilia. Am. J. Physiol.Gastrointest. Liver Physiol. 291:G500–9

166. Collin SP, Barry Collin H. 2004. Primary cilia in vertebrate corneal endothelialcells. Cell Biol. Int. 28:125–30

167. Gallagher BC. 1980. Primary cilia of the corneal endothelium. Am. J. Anat.159:475–84

168. Bystrevskaia VB, Antonov AS, Perov NA. 1987. Primary cilium in the endothe-lial cells of the human aorta. Dokl. Akad. Nauk. SSSR 297:1233–36

169. Iomini C, Tejada K, Mo W, Vaananen H, Piperno G. 2004. Primary cilia ofhuman endothelial cells disassemble under laminar shear stress. J. Cell Biol.164:811–17

170. Van der Heiden K, Groenendijk BC, Hierck BP, Hogers B, Koerten HK, et al.2006. Monocilia on chicken embryonic endocardium in low shear stress areas.Dev. Dyn. 235:19–28

171. Haust MD. 1987. Endothelial cilia in human aortic atherosclerotic lesions. Vir-chow? Arch. A Pathol. Anat. Histopathol. 410:317–26

172. Tobe K, Tsuchiya T, Itoshima T, Nagashima H, Kobayashi T. 1985. Electronmicroscopy of fat-storing cells in liver diseases with special reference to ciliaand cytoplasmic cholesterol crystals. Arch. Histol. Jpn. 48:435–41

173. Tritton DJ. 1977. Physical Fluid Mechanics. New York: Van Nostrand Reinhold174. Schwartz EA, Leonard ML, Bizios R, Bowser SS. 1997. Analysis and modeling

of the primary cilium bending response to fluid shear. Am. J. Physiol. RenalPhysiol. 272:F132–38

175. Gittes F, Mickey B, Nettleton J, Howard J. 1993. Flexural rigidity of micro-tubules and actin filaments measured from thermal fluctuations in shape. J. CellBiol. 120:923–34

176. Liu W, Xu S, Woda C, Kim P, Weinbaum S, Satlin LM. 2003. Effect of flowand stretch on the [Ca2+]i response of principal and intercalated cells in corticalcollecting duct. Am. J. Physiol. Renal Physiol. 285:F998–1012

177. Praetorius HA, Frokiaer J, Nielsen S, Spring KR. 2003. Bending the primary cil-ium opens Ca2+-sensitive intermediate-conductance K+ channels in MDCKcells. J. Membr. Biol. 191:193–200

178. McGlashan SR, Jensen CG, Poole CA. 2006. Localization of extracellularmatrix receptors on the chondrocyte primary cilium. J. Histochem. Cytochem.54(9):1005–14

179. Mijailovich SM, Kojic M, Zivkovic M, Fabry B, Fredberg JJ. 2002. A finite ele-ment model of cell deformation during magnetic bead twisting. J. Appl. Physiol.93:1429–36

180. Nagayama K, Nagano Y, Sato M, Matsumoto T. 2006. Effect of actin filamentdistribution on tensile properties of smooth muscle cells obtained from ratthoracic aortas. J. Biomech. 39:293–301


Ann

u. R

ev. B

iom

ed. E

ng. 2

007.

9:1-

34. D

ownl

oade

d fr

om a

rjou

rnal

s.an

nual

revi

ews.

org

by C

APE

S on

05/

26/0

9. F

or p

erso

nal u

se o

nly.

ANRV317-BE09-01 ARI 7 June 2007 15:9

181. Byfield FJ, Aranda-Espinoza H, Romanenko VG, Rothblat GH, Levitan I. 2004.Cholesterol depletion increases membrane stiffness of aortic endothelial cells.Biophys. J. 87:3336–43

182. Caille N, Thoumine O, Tardy Y, Meister JJ. 2002. Contribution of the nucleusto the mechanical properties of endothelial cells. J. Biomech. 35:177–87

183. Sugawara M, Ishida Y, Wada H. 2004. Mechanical properties of sensory andsupporting cells in the organ of Corti of the guinea pig cochlea—study by atomicforce microscopy. Hear Res. 192:57–64

184. Sugawara M, Ishida Y, Wada H. 2002. Local mechanical properties of guineapig outer hair cells measured by atomic force microscopy. Hear Res. 174:222–29

185. Mahaffy RE, Park S, Gerde E, Kas J, Shih CK. 2004. Quantitative analysisof the viscoelastic properties of thin regions of fibroblasts using atomic forcemicroscopy. Biophys. J. 86:1777–93

186. Koay EJ, Shieh AC, Athanasiou KA. 2003. Creep indentation of single cells.J. Biomech. Eng. 125:334–41

187. Hochmuth RM. 2000. Micropipette aspiration of living cells. J. Biomech. 33:15–22

188. Roca-Cusachs P, Almendros I, Sunyer R, Gavara N, Farre R, Navajas D. 2006.Rheology of passive and adhesion-activated neutrophils probed by atomic forcemicroscopy. Biophys. J. 91(9):3508–18

189. Peeters EA, Oomens CW, Bouten CV, Bader DL, Baaijens FP. 2005. Vis-coelastic properties of single attached cells under compression. J. Biomech. Eng.127:237–43

190. Laurent VM, Fodil R, Canadas P, Fereol S, Louis B, et al. 2003. Partitioning ofcortical and deep cytoskeleton responses from transient magnetic bead twisting.Ann. Biomed. Eng. 31:1263–78

191. Lekka M, Laidler P, Gil D, Lekki J, Stachura Z, Hrynkiewicz AZ. 1999. Elas-ticity of normal and cancerous human bladder cells studied by scanning forcemicroscopy. Eur. Biophys. J. 28:312–16

192. Takai E, Costa KD, Shaheen A, Hung CT, Guo XE. 2005. Osteoblast elasticmodulus measured by atomic force microscopy is substrate dependent. Ann.Biomed. Eng. 33:963–71

193. Ananthakrishnan R, Guck J, Wottawah F, Schinkinger S, Lincoln B, et al. 2006.Quantifying the contribution of actin networks to the elastic strength of fibrob-lasts. J. Theor. Biol. 242:502–16

194. Park S, Koch D, Cardenas R, Kas J, Shih CK. 2005. Cell motility and localviscoelasticity of fibroblasts. Biophys. J. 89:4330–42

195. Coughlin MF, Puig-de-Morales M, Bursac P, Mellema M, Millet E, Fredberg JJ.2006. Filamin-a and rheological properties of cultured melanoma cells. Biophys.J. 90:2199–205

196. Lee JS, Panorchan P, Hale CM, Khatau SB, Kole TP, et al. 2006. Ballisticintracellular nanorheology reveals ROCK-hard cytoplasmic stiffening responseto fluid flow. J. Cell Sci. 119:1760–68

197. Desprat N, Richert A, Simeon J, Asnacios A. 2005. Creep function of a singleliving cell. Biophys. J. 88:2224–33


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Annual Reviewof BiomedicalEngineering

Volume 9, 2007Contents

Cell Mechanics: Integrating Cell Responses to Mechanical StimuliPaul A. Janmey and Christopher A. McCulloch � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � 1

Engineering Approaches to BiomanipulationJaydev P. Desai, Anand Pillarisetti, and Ari D. Brooks � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � 35

Forensic Injury BiomechanicsWilson C. Hayes, Mark S. Erickson, and Erik D. Power � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � 55

Genetic Engineering for Skeletal Regenerative MedicineCharles A. Gersbach, Jennifer E. Phillips, and Andrés J. García � � � � � � � � � � � � � � � � � � � � � � 87

The Structure and Function of the Endothelial Glycocalyx LayerSheldon Weinbaum, John M. Tarbell, and Edward R. Damiano � � � � � � � � � � � � � � � � � � � � � �121

Fluid-Structure Interaction Analyses of Stented Abdominal AorticAneurysmsC. Kleinstreuer, Z. Li, and M.A. Farber � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �169

Analysis of Time-Series Gene Expression Data: Methods, Challenges,and OpportunitiesI.P. Androulakis, E. Yang, and R.R. Almon � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �205

Interstitial Flow and Its Effects in Soft TissuesMelody A. Swartz and Mark E. Fleury � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �229

Nanotechnology Applications in CancerShuming Nie, Yun Xing, Gloria J. Kim, and Jonathan W. Simons � � � � � � � � � � � � � � � � � �257

SNP Genotyping: Technologies and Biomedical ApplicationsSobin Kim and Ashish Misra � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �289

Current State of Imaging Protein-Protein Interactions In Vivo withGenetically Encoded ReportersVictor Villalobos, Snehal Naik, and David Piwnica-Worms � � � � � � � � � � � � � � � � � � � � � � � � � � �321

Magnetic Resonance–Compatible Robotic and Mechatronics Systemsfor Image-Guided Interventions and Rehabilitation: A Review StudyNikolaos V. Tsekos, Azadeh Khanicheh, Eftychios Christoforou,and Constantinos Mavroidis � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �351

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AR317-FM ARI 7 June 2007 19:9

SQUID-Detected Magnetic Resonance Imaging in Microtesla FieldsJohn Clarke, Michael Hatridge, and Michael Moßle � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �389

Ultrasound Microbubble Contrast Agents: Fundamentals andApplication to Gene and Drug DeliveryKatherine Ferrara, Rachel Pollard, and Mark Borden � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �415

Acoustic Detection of Coronary Artery DiseaseJohn Semmlow and Ketaki Rahalkar � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �449

Computational Anthropomorphic Models of the Human Anatomy:The Path to Realistic Monte Carlo Modeling in Radiological SciencesHabib Zaidi and Xie George Xu � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �471

Breast CTStephen J. Glick � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �501

Noninvasive Human Brain StimulationTimothy Wagner, Antoni Valero-Cabre, and Alvaro Pascual-Leone � � � � � � � � � � � � � � � � � � �527

Design of Health Care Technologies for the Developing WorldRobert A. Malkin � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �567

Indexes

Cumulative Index of Contributing Authors, Volumes 1–9 � � � � � � � � � � � � � � � � � � � � � � � � � � �589

Cumulative Index of Chapter Titles, Volumes 1–9 � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � � �593

Errata

An online log of corrections to Annual Review of Biomedical Engineering chapters(if any, 1977 to the present) may be found at http://bioeng.annualreviews.org/

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