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Abstract

Cells process various mechanical cues in the microenvironment to self-organize into high-order architectures during tissue morphogenesis. Impairment of morphogenic processes is the underlying cause of many diseases; as such, understanding the regulatory mechanisms associated with these processes will form the foundation for the development of innovative approaches in cell therapy and tissue engineering. Nevertheless, little is known about how cells collectively respond to mechanical cues in the microenvironment, such as global geometric guidance, local cell-cell interactions, and other physicochemical factors, for the emergence of the structural hierarchy across multiple length scales. To elucidate the mechanoregulation of tissue morphogenesis, numerous approaches based on biochemical, biomaterial, and biophysical techniques have been developed in the past decades. In this review, we summarize techniques and approaches for probing the mechanoregulation of tissue morphogenesis and illustrate their applications in vasculature development. The potential and limitations of these methods are also discussed with a view toward the investigation of a wide spectrum of tissue morphogenic processes.

Keywords

automated biology microtechnology nanotechnology

Introduction

Tissue morphogenesis is a fundamental multicellular activity that is essential for various developmental and regenerative processes. Vasculogenesis, for instance, leads to de novo blood vessel formation from mesodermal precursors overlying the endoderm.¹ Organs of ectodermal origin, on the other hand, are vascularized by angiogenesis—the outgrowth of new capillaries from the existing vasculature.² The formation of new blood vessels also contributes to numerous malignant, ischemic, inflammatory, infectious, and immune disorders.^3,4 Particularly, tissue engineering, a rapidly emerging field in regenerative medicine, has great potential to restore, maintain, or improve tissue functions. Skin and cartilage tissue engineering, for example, has gained significant success because of the relative simplicity of the tissue architecture. Other tissues with extensive vasculature and heterogeneous cell arrangements, however, present significant hurdles in creating functional tissues that mimic their physiological counterparts.^5,6 A fundamental understanding of the mechanisms and regulatory factors involved is required for creating functional tissue constructs for regenerative medicine.

The chemical basis of biological pattern formation and tissue morphogenesis is best understood in the reaction-diffusion model, which is also referred to as the activator-inhibitor system or the Turing pattern.⁷ Moreover, the theoretical model describes the roles of autocatalytic reactions and lateral inhibition in tissue morphogenesis and has been applied to explain a wide spectrum of morphogenic processes.^8–11 In addition to the reaction and diffusion of morphogens, physical factors are also important throughout the developmental process. The regulatory role of cell mechanics (e.g., cell contractility, intercellular tension, and cell-matrix mechanical interactions) is increasingly recognized in tissue morphogenesis.¹² As an example, geometric confinement has been demonstrated to regulate capillary network topology via cell-matrix mechanical interactions.¹³ Tissue deformation has been shown to modulate vascular endothelial growth factor gradients and endothelial cell proliferation in deformable tissue constructs, coupling biochemical and mechanical tissue regulation.¹⁴ Cell traction force has been demonstrated to control capillary network formation in vitro and in vivo by applying a Rho inhibitor and modulating extracellular matrix (ECM) elasticity.¹⁵ Alternation of the ECM composition also revealed the importance of endothelial cell traction force in network stabilization.¹⁶ These studies highlight the mechanical basis of tissue morphogenesis and encourage the development of novel mechanoregulation techniques.

Cells in tissues interact with their physical environment and generate endogenous contractile forces via multiple feedback mechanisms.^17–19 The cell traction forces are coordinated by the cytoskeleton dynamics as well as cell-cell and cell-ECM interactions. Advances in biochemistry, material science, microfabrication, and nanotechnology have created new opportunities in modulating these interactions and probing the mechanoregulation of tissue morphogenesis systematically.^20–24 In this article, we summarize advances in techniques and approaches for studying the roles of mechanical factors in tissue morphogenesis. First, pharmacological and biochemical approaches for perturbing the cytoskeletal structures and cell adhesion molecules are discussed. Second, biomaterial and microfabrication techniques for modulating the cell-matrix mechanical interactions are presented. External physical perturbations with mechanical, optical, magnetic, and fluidic techniques are then reviewed. Examples of the development of vascular and other tissues are described to illustrate the applicability of these techniques. Finally, the potential and limitations of these techniques for probing the mechanoregulation of tissue morphogenesis are discussed.

Biochemical Approaches for Mechanical Perturbation

Actin-Targeting Reagents

Cell traction forces and intercellular mechanical interactions can be modulated by reagents that target various cellular components such as actin filaments, microtubules, actomyosin, and cell junctions. The chemical inhibitors, RNA interference, forced expression, as well as genetic mutations of cellular components have been used to alter the mechanical properties of cells ( Fig. 1 ). It is well known that the actin cytoskeleton plays a major role in mechanical support, generation of cell traction force, and transmission of exogenous and endogenous cellular stress; therefore, actin-targeting drugs possess a significant capability to perturb cell mechanics. For example, cytochalasins, which are F-actin (filamentous) depolymerization reagents, bind to the barbed end of F-actin and prevent actin monomer polymerization. With cytochalasin D treatment, the contractile force and dynamic stiffness of endothelial cells are reduced significantly, whereas the intracellular structural damping remains unaffected.²⁵ In angiogenesis studies, endothelial cell tube formation is also inhibited by cytochalasin D.²⁶ Interestingly, at low concentrations of cytochalasin D, changes in the actin cytoskeleton are undetectable by fluorescence microscopy, despite significant changes in the mechanical properties of the cells.²⁷ Cytochalasin E, similar to cytochalasin D, inhibits endothelial proliferation and angiogenesis without disrupting actin stress fibers.²⁸

Figure 1.

Biochemical approaches for perturbing cell mechanics.

Latrunculin, a class of actin depolymerizing agents,^29,30 is more potent than cytochalasins. Latrunculin B binds to G-actin (globular) and inhibits nucleotide exchange on actin, while reducing cell stiffness by approximately 50%.³¹ Low concentrations of Latrunculin B preferentially inhibit F-actin polymerization in filopodia.³² Y-27632, a synthetic pyridine derivative that inhibits the Rho kinases (ROCK I and ROCK II), affects various downstream events including F-actin depolymerization and inhibition of myosin activity. As a contraction inhibitor, it relaxes contraction in smooth muscle cells and trabecular meshworks.³³ Treatment with Y-27632 abrogated vascular endothelial growth factor (VEGF)–induced softening of human umbilical vein endothelial cells in three-dimensional (3D) matrixes³⁴ and inhibited angiogenesis in vivo without observable effects on preexisting vessels.³⁵ In addition to chemically regulating the actin cytoskeleton, RNA interference of actin can also be applied to modulate mechanical forces. For example, the interference of smooth muscle actin (α-SMA) attenuated transforming growth factor β (TGF-β)–induced cell traction force in ﬁbroblasts, and a linear relationship was also observed between the α-SMA protein expression level and the magnitude of cell traction force.³⁶

Microtubule-Targeting Molecules

Microtubule-targeting drugs represent another group of potent agents for modulating cell mechanics; however, the mechanical effects of microtubule-targeting drugs are often ultimately related to the actin cytoskeleton.³⁷ Nocodazole, a synthetic benzimidazole that destabilizes microtubules, disrupts the cellular balance between actin and microtubule networks and indirectly activates the actin cytoskeleton to improve force generation.³⁸ In addition, nocodazole exhibits anti-angiogenic effects in vitro along with potent inhibitory effects on endothelial growth.³⁹ Taxol (paclitaxel), an agent that stabilizes microtubules, facilitates microtubule polymerization and increases cell stiffness and viscosity.⁴⁰ Taxol has been actively investigated for its anti-angiogenic effects both in vitro and in vivo.⁴¹ Interestingly, taxol, in low concentrations, has been shown to inhibit angiogenesis in vitro without affecting microtubule assembly.⁴²

Drugs Targeting Actin-Myosin Interactions

The modulation of cell mechanics in physiological and pathological processes is also achieved by controlling actin-myosin interactions. Myosins are a family of ATP-dependent motor molecules responsible for cell contraction and motility in both muscle and nonmuscle cells. For instance, 2,3-butanedione monoxime, an inhibitor of the ATPase activity of myosin, depressed the contractile characteristics of muscle cells⁴³ and decreased the stiffness of cardiomyocytes.⁴⁴ ML-7 and ML-9, agents that bind to myosin light chain kinase competitively with ATP, can also be used to reduce cell stiffness.^45,46 Blebbistatin, a pharmacological inhibitor that restricts actin-myosin interaction by lowering myosin’s affinity for actin, reduces contractile force and cadherin adhesion.⁴⁷

RNA interference can be applied to inhibit actomyosin interactions and angiogenesis.⁴⁸ However, myosin inhibition through RNA interference also influences the mechanical properties of the cells. Myosin IIA small interfering RNA has been demonstrated to markedly reduce the contraction force of fibroblast cells within fibrins.³⁸ The inhibitor of an unconventional myosin, Myo1G, was also shown to decrease the elasticity of Jurkat cells significantly.⁴⁹

Targeting Intermediate Filaments

Another major component of the cytoskeletal system is the intermediate filament network. Intermediate filaments play essential roles in providing mechanical and structural integrity for cells, including regulating cellular tension development.⁵⁰ Multiple types of intermediate filament proteins have been identified. Among them, vimentin is one of the most widely distributed intermediate filament proteins. Vimentin intermediate filaments support cellular membranes, fix the position of some organelles, and transmit membrane receptor signals to the nucleus. Stimulation of tracheal smooth muscle strips with acetylcholine induced the increase in the ratio of soluble to insoluble vimentin in association with force development.⁵¹ Treatment of muscle tissues with vimentin RNA interference attenuated force development in response to acetylcholine or KCl depolarization, as well as lower passive tension; the latter may be associated with the impairment of desmosomes.⁵²

Targeting Focal Adhesions and Cell Junctions

Cell mechanics may also be modulated by targeting cell-matrix and cell-cell interactions with genetic manipulation. Vinculin, which couples integrins or cadherins to the actin cytoskeleton at focal adhesions and adherens junctions, has been frequently used for assessing cell mechanical measurements. Cell contractile force generation is reduced when vinculin is absent or enhanced when vinculin is up-regulated.⁵³ Depleting the paxillin-vinculin interaction by substituting endogenous paxillin with a mutant reduced the total traction force of mouse embryonic fibroblasts.⁵⁴ Overexpression of the vinculin binding domain of αE-catenin decreased E-cadherin–mediated adhesion strength.⁵⁵ Down-regulation of talin I, an adaptor protein that links integrins to actin at the adhesion complex, decreased cellular force generation.⁵⁶ Depletion of α-actinin, which links integrins with actin, enhanced initial force generation and prevented adhesion mutation in subsequent steps.⁵⁷ Overexpression of actin-binding protein caldesmon blocked cell contractility and interfered with focal adhesion formation.⁵⁸ Chemical inhibitions of focal adhesion kinase with specific inhibitor or broad spectrum inhibitor have been shown to increase traction forces.⁵⁹ Activation of integrins with integrin-activating antibody inhibited the contractile force of muscle cells. Similar effects can be achieved by treatment with a synthetic integrin-binding peptide, whereas integrin function-blocking antibodies reversed the effect of the peptide on contractile force.⁶⁰

Targeting Signaling Pathways That Regulate Cell Mechanics

The manipulation of several signaling pathways can directly or indirectly lead to changes in cell mechanics. RNA interference of transcription factor JunB led to the inhibition of cell contractility under both basal and TGFβ1-stimulated conditions.⁶¹ Treatment of DNA binding protein HMGB1 caused a TLR4-dependent increase in traction force, accounting for the downstream inhibition of enterocyte migration.⁶² Silencing of the function-unknown gene VPS13A attenuated the F-actin network and reduced the stiffness of endothelial cells.⁶³ Silencing integrin-linked kinase enhanced vascular smooth muscle cell contraction under a puling force.⁶⁴ Inhibition of soluble adenylyl cyclase results in significant endothelial cell softening.⁶⁵ However, for applying biochemical approaches in studying cell mechanoregulation, achieving a pure mechanical perturbation is still a challenge because of the complex signaling network. Analyzing the results due to the force feedback–sensing, as well as force feedback–generating, mechanisms in cells is also demanding.

Biomaterial Approaches for Modulating the Microenvironment

ECM and Substrate Properties

Cell-matrix interactions can be modulated by adjusting the matrix properties (e.g., stiffness) as well as by altering ECM geometry and/or topography ( Table 1 ). The stiffness of the matrix can be controlled by the ECM density. Increasing the density of fibrin from 2.5 to 10 mg/mL resulted in an approximately sevenfold increase in matrix stiffness.⁶⁶ Fibrin gel has been used to study 3D capillary morphogenesis, with stiff gels found to reduce the formation of capillary networks.^16,67 Similarly, the extent of capillary-like network formation could be tuned by adjusting the density of a self-assembling peptide gel.⁶⁸ Despite the simplicity of this approach, adjusting the matrix stiffness via the gel density will also affect the ligand density, which could introduce ambiguity into the results.

Table 1.

Common Gel-Based Materials for Studying Cell-Matrix Mechanical Interactions.

ECM Type	ECM Component	Ligand Density	Stiffness Range	Model and Parameters	Effects
Protein	Fibrin^66,67	2.5–10 mg/mL	1.3–9.0 kPa	HUVEC coated beads	Decreased
Hydrogel				Total length and number of sprouts	3-fold
	Glycated collagen⁶⁹	1.5 mg/mL	175–515 Pa	BAEC spheroids	Decreased
				Same as above	2- and 1.5-fold
Half synthetic	PA/collagen⁷⁰	0.1 mg/mL	0.2–10 kPa	Capillary-like structures	Form on ECM
Hydrogel					Stiffness <1 kPa
	Collagen/PA/glass	1.6 mg/mL	0.3–14 kPa	Pseudopodial branching	Inhibited
	Sandwich⁷¹				Fewer branches
Fully synthetic	Peptide⁶⁸	1–3 mg/mL	46–735 Pa	Capillary-like network	Decreased
Hydrogel				Average structure size	474–60 µm
	Degradable	10 mg/mL	3.17–0.62 kPa	Vascular sprout	Stiffness
	PEGDA⁷²			Direction	Dependent

BAEC, bovine aortic endothelial cell; ECM, extracellular matrix; HUVEC, human umbilical vein endothelial cell; PA, polyacrylamide; PEGDA, polyethylene glycol diacrylate hydrogel.

Glycation of ECM proteins (e.g., type I collagen) has been employed to increase ECM stiffness with constant ligand density. More specifically, collagen gels were incubated in glucose-6-phopshate for 5 and 8 d to achieve different levels of glycation and ECM stiffness. Using collagen glycation, sprouting angiogenesis was shown to be delayed in stiff gels in a co-culture (endothelial cells and smooth muscle cells) sprouting model.⁷³ Alternatively, collagen solutions could be mixed with ribose to form glycated collagen solutions with increased concentrations of ribose and stiffness (~175 to ~730 Pa), leading to an increased number and length of sprouts from endothelial cell spheroids.⁶⁹ However, the time for complete glycation is long and the range of comprehensive moduli is small in comparison to the relevant physiological range (hundreds of kPa). Biological inert synthetic polymers have also been developed for achieving tunable gel stiffness with constant ligand density.^74,75

Polyacrylamide (PA) hydrogels may achieve a large range of ECM stiffness by simply altering the relative concentrations of monomer (acrylamide) and cross-linker (bis-acrylamide).⁷⁶ However, it is not compatible with cell culture directly because of potential toxicities as well as the lack of cell adhesion capability. Proper PA hydrogel modification is required for investigating the effects of mechanical interactions at the cell-substrate interface. Endothelial cells have been shown to self-organize into capillary-like structures on compliant PA gels derivatized with type I collagen or functionalized with Arg-Gly-Asp (RGD) peptides.^70,77 A collagen/PA/glass sandwich gel that enabled 3D cell culture was also shown to manipulate substrate stiffness, revealing that high ECM stiffness inhibits endothelial pseudopodial branch initiation.⁷¹

Micropost arrays have also been used for modulating substrate rigidity, by taking advantage of microfabrication technology. The micropost arrays have been primarily used for studying cell mechanical interactions on 2D substrates and can be modified for studying 3D microtissues.⁷⁸ The deformability of the micropost can be modified by adjusting the post dimensions (e.g., height). Micropost arrays allow independent control of surface properties, decoupling substrate rigidity from cell adhesion.^79,80 Furthermore, for compliant microposts (e.g., polydimethylsiloxane [PDMS]), cell traction forces can be determined by measuring the micropost displacement. It has been reported that endothelial cells showed enhanced elongation and alignment on PDMS microposts compared with stiff SiO₂ microposts with similar topographical features.⁸¹ However, the effects of ECM roughness and topology cannot be ignored and may limit micropost applications on stiffness-associated cell research.

Dynamic tuning of local ECM stiffness was demonstrated with polyethylene glycol (PEG)–based photodegradable hydrogels, in which cells exhibited a rounded morphology initially and started spreading during irradiation-induced matrix stiffening.⁸² In addition, light-mediated sequential cross-linking was applied for dynamic matrix stiffening (e.g., 3–30 kPa), which mimicked the dynamic nature of tissue development, wound healing, and pathogenesis.⁸³ Increased cell areas and traction forces were observed in adhered human mesenchymal stem cells over a time scale of hours as the substrates were stiffened.

PA hydrogels with a photo initiator have been used for manipulating spatial control of the matrix stiffness. Stiffness gradients were generated by progressively uncovering the gel solution with an opaque mask from a noncollimated ultraviolet lamp.⁸⁴ Alternatively, a matrix metalloproteinase (MMP)–sensitivity PEG diacrylate hydrogel (PEGDA) was used to generate stiffness gradients via matrix degradation.⁷² The stiffness gradients were capable of directing 3D vascular sprout formation using a co-culture angiogenesis model. In addition, magnetic beads could be embedded in collagen gels via bio-conjugation to alter the local stiffness of the ECM at the presence of an external magnetic field.⁸⁵ The magnetic force could increase the apparent stiffness of the ECM, and magnetic force gradients could induce ECM stiffness gradients, which can affect endothelial cell behaviors during angiogenesis.

Geometric Control

ECM geometries and physical confinements created by lithography and other microfabrication techniques can also modulate tissue morphogenesis.^86,87 Photolithographic techniques were applied for micropatterned PEGDA with adhesive ligands to regulate and guide endothelial morphogenesis.⁸⁸ Endothelial cells did not assemble into cordlike structures on the stripe that was larger than 50 µm in width, highlighting the importance of geometric control of endothelial morphogenesis. Geometric control of endothelial cord formation was also demonstrated by culturing cells in microchannels that were filled with collagen gels.⁸⁹ The microchannel confinement is also used for directing the development of branches in tube formation, which may enable the production of complex capillary architectures. More recently, endothelial cells cultured on narrow (10 and 50 µm) micropatterned angiogenic Ser−Val−Val− Tyr−Gly−Leu−Arg (SVVYGLR) peptides demonstrated restricted spreading, with orientation and migration directionally guided and regulated.⁹⁰ Microwells with arbitrary shapes (e.g., star, square, and triangle) were shown to module capillary topology via cell-matrix mechanical interactions.¹³ Endothelial cells can form denser networks on acute angle corners than those on reflex angle corners in a star-shaped ECM structure. The geometric control of tissue morphogenesis has significant implications in microfabrication-based tissue models and scaffold design for tissue engineering.

ECM topography and fiber alignment can also affect cell behaviors, such as the orientation of actin filaments and focal adhesions, proliferation, and migration.^91–93 In particular, endothelial progenitor cells on nanotopographic substrates (600-nm-wide ridge and groove) exhibited enhanced alignment, organization, and capillary tube formation.⁹⁴ With the improvement and development of micro- and nanofabrication technologies in the future, mimicking in vivo ECM with specific topology for investigating tissue morphogenesis will be possible.

Biophysical Approaches for Probing Tissue Morphogenesis

Mechanical Perturbation

Mechanical perturbation influences various cell functions, including proliferation, migration, differentiation, and ECM remodeling ( Table 2 ). Tissue deformation that occurs naturally in muscular and cardiovascular systems regulates various tissue morphogenic processes.^95,96 Mechanical stretching of rat microvascular endothelial cells enhanced the production of MMP-2, MT1-MMP, HIF-1α, HIF-2α, and VEGF at both the transcriptional and translational levels.⁹⁷ Stretching also induced an increase in intracellular Ca²⁺ concentration and traction forces of NIH-3T3 fibroblasts via stretch-activated ion channels.⁹⁸ In addition, physical stimulation with a mechanical probe may also be applied to create local mechanical stress and injury to individual cells in the tissue.^99–101 A mechanical probe with comb-drive (capacitive) force measurement, for instance, has been used to study mechanical stimulation–induced intercellular calcium communication in microengineered endothelial networks.^102,103 Endothelial cells collectively regulated their calcium levels against a large range of probing forces and repeated stimulations. Moreover, endothelial cells on elastic gels were shown to have active calcium responses when stimulated with a vibrating probe.¹⁰⁴ The mechanical stimulation caused deformations over a distance from the probe, and the cells sensed the local stimulation.

Table 2.

Biomechanical Effects on Tissue Morphogenesis.

Mechanical Perturbations	Tissue or Cell Type	Approach	Tissue or Cell Response
Mechanical stretch	Rat microvascular endothelial cells, Sprague-Dawley rats	Overload rat extensor digitorum longus muscles	Increased capillary number^105,106
	Fibroblast	Stretch elastic gel	Increased leading edge proliferation, proliferation rate^100,107
Optical technique	HUVECs	Manipulate fibronectin-coated beads	Rapid distal Src activation and slower directional wave propagation of Src activation¹⁰⁸
	HMVECs	Apply tensile and compressive force	Increased capillary growth¹⁰⁹
	ECs, Drosophila embryos	Laser ablation	Increase cell migration, neural regeneration, morphgenic movement^110–112
Magnetic technique	HUVECs	Adjust magnetic fields	Accelerate cell migration, regulate migration direction¹¹³
	Smooth muscle cells	Uniform-gradient magnetic manipulator	3D cellular deformation¹¹⁴
	Drosophila embryos	Manipulate magnetic particles	Compression of stomodeal cells¹¹⁵
	Microposts	Magnetic microposts	Increase in local focal adhesion size¹¹⁶
Shear stress	Rabbit ear chamber, rat microvascular endothelial cells, Sprague-Dawley rats	Parallel plate flow chamber	Increased capillary density, capillary number, and capillary growth rate^105,106,117
	BAEC, HUVECs, bovine pulmonary microvascular endothelial cells	Parallel-plate flow chamber	Enhanced sprouting,¹¹⁸ directional assembly¹¹⁹ directs cell sprouting¹²⁰

3D, three-dimensional; BAEC, bovine aortic endothelial cell; EC, endothelial cell; HMVEC, human microvascular endothelial cell; HUVEC, human umbilical vein endothelial cell.

Optical Techniques

Optical techniques can generate mechanical perturbations with subcellular resolution. As an example, Src activities in single endothelial cells could be activated mechanically by laser tweezers with fibronectin-coated beads.¹²¹ Optical tweezers in a microfluidic system generated tensile and compressive forces for studying focal adhesion recruitment in endothelial cells.¹²² In particular, mechanical force was applied to the ECM-integrin-cytoskeleton linkage by trapping and manipulating functionalized beads. In addition to laser tweezers, photoablation of cells and subcellular structures with lasers represented another powerful approach in modulating cell mechanics with subcellular resolution.¹⁰⁸ Cell ablation may be applied to disrupt the mechanical interactions between cells and perturb specialized cells in the morphogenic process. For disrupting the cytoskeleton, fluorescent proteins, such as actin-GFP, were used to visualize the cytoskeletal structure, and lasers were applied to ablate or disrupt the cytoskeleton. Femtosecond lasers with multiphoton absorption enabled fine resolution (<300 nm) and minimized damage to surrounding structures. The photoablation technique has been used for investigating various tissue morphogenic processes, such as collective cell migration,¹⁰⁹ neural regeneration,¹⁰⁸ and morphogenic movements in Drosophila embryos.¹¹⁰

Magnetic Technique

Magnetic tweezers are employed to apply mechanical forces to living cells and tissues via external magnetic field gradients.¹¹¹ Intracellular forces on endothelial cells are generated under different magnetic fields after cellular uptake of superparamagnetic iron oxide nanoparticles. The intracellular magnetic force accelerates cell migration and regulates cell migratory direction via adjusting magnetic fields and increasing the cell-free culture space.¹¹² Moreover, 3D cellular deformation was studied by applying uniform forces to a large cell population via a uniform-gradient magnetic manipulator.¹¹³ Magnetic particles were injected into Drosophila embryos for in vivo studies of tissue deformation. The mechanical compression of stomodeal cells in embryo mimicked the physiological deformation experienced by stomodeal cells due to germ band extension at the onset of gastrulation, which could up-regulate Twist expression in the stomodeal primordium.¹¹⁴ Magnetic microposts were also employed to exert a step force that led to an increase in local focal adhesion size at the site of application but not at nearby nonmagnetic posts.¹¹⁵

Shear Stress

Endothelial cells are subjected to shear stress generated by blood flow. Shear stress can modulate cell migration, proliferation, gene expression, and morphology of endothelial cells.^116,123 Thus, wall shear stress can be generated to investigate the effects of dynamic shear on the development and remodeling of the vasculature. A parallel plate flow chamber may be used to study the effects of shear stress on capillary-like tube formation. Endothelial cells on matrigel experienced high shear stress, resulting in aligned tubular structures along with the direction of fluid flow. Shear stress–dependent regulation was, at least partially, induced by VEGF expression.^{105,117,120,124} Fluid forces generated by shear stress may also attenuate endothelial cell sprouting in a nitric oxide–dependent manner. Interstitial flow directs cell sprouting and cell morphology and therefore regulates capillary tube formation. Moreover, fluidic shear stress (3 dyn/cm²) enhanced the sprouting of bovine pulmonary microvascular endothelial cells in a 3D culture model.¹²⁰ In vivo, the effects of fluid shear stress in capillary vessel growth were first investigated in rat skeletal muscle and rabbit ear chamber.^106,118 Externally applied wall shear stress was shown to stimulate angiogenesis via induction of VEGF expression by HIF-1α via the PI3K-dependent Akt phosphorylation pathway.

Microfluidic technology provides powerful tools for investigating the effects of fluidic shear stress on tissue morphogenesis in 2D and 3D models.¹²⁵ Fluid shear can be applied to perturb diffusible gradients in the microenvironment, thereby isolating biomechanical mechanisms from other biochemical mechanisms (e.g., reaction-diffusion). For instance, microfluidics was used to identify autocatalytic alignment feedback in the long-range organization of myotube development.¹²⁶ While diffusible factors are essential in the alignment process, recirculation of the media results in normal cell fusion and long-range alignment, eliminating diffusible factors from the cell alignment mechanism. Furthermore, a microfluidic wound-healing assay was developed to wound the monolayer and test the effects of shear stress on cell migration.¹²⁷

Discussion

This review summarizes biochemical, biomaterial, and biophysical approaches for probing the regulatory roles of mechanical force in tissue morphogenesis. Several global perturbation techniques, such as modulation of ECM properties, fluid shear, and mechanical stretching, have been used extensively because of their physiological and translational relevance. These techniques are particularly useful in identifying the involvement of mechanical forces in morphogenic processes. The overall tissue architectures (e.g., orientation and network density) can be perturbed to shed light on the morphogenic process. Advances in technology have also enabled local perturbation techniques, such as single-cell probes, laser tweezers, microfluidics, and photoablation, for investigating cell-cell communication mechanisms. These techniques open new possibilities in elucidating morphogenic mechanisms by perturbing the local mechanical force distribution and observing its effects on the tissue architecture. In particular, these local perturbation approaches can be used for exploring unknown morphogenic processes, testing new hypotheses, and studying cell-cell organization. Furthermore, biochemical reagents provide invaluable insights into the molecular mechanisms involved in the mechanoregulation processes. However, biochemical reagents may disturb multiple signaling pathways and trigger nonspecific effects, which complicates the interpretation of results. Multiple reagents and approaches should be incorporated to confirm that the observation is indeed mechanically induced.

Cells and tissues represent complex networks of mechanically responsive systems with multiple hierarchical levels. Cells in tissues collectively sense and adapt to the physical microenvironment during development and regeneration. Intra- and intercellular forces are dynamically regulated through multiple cellular feedback mechanisms. While it is important to understand the molecular mechanosensing mechanism, it is challenging, if not impossible, to isolate individual factors one from the other. This represents a hurdle in understanding and interpreting study results and in elucidating mechanical regulatory mechanisms. Another challenge in understanding tissues morphogenesis is to identify the emerging behaviors that drive complex tissue architectures. Inhibiting a morphogenic process does not necessarily provide useful information in the conceptual understanding of the emerging behaviors. Careful experimental designs with multiple techniques are often required to study collective cell behavior. In addition to mechanical perturbation techniques, novel biochemical and mechanical techniques (e.g., intracellular probes^100,128 and traction force microscopy^129,130) should be developed and used to study morphogenic processes. Another key challenge in studying the emergent complexity of biological tissues is the difficulty in understanding the high-order structures and functions resulting from local interactions of individual cells. These emergent properties can often be counterintuitive and cannot be understood by simply extrapolating interaction between a few cells. The complex interplays between global microenvironmental cues and local cell-cell interactions further complicate understanding emergent cell behavior. A complex systems framework that incorporates biomanufacturing, microfluidics, advanced materials, biosensors, and computational modeling is ultimately required to fully understand the mechanoregulation of tissue morphogenesis. As we move forward in this field to understand this complexity, not only will basic cellular and tissue processes become clear, this insight may additionally be used in a biomanipulative way to develop novel therapeutic strategies to tackle complex disease processes that involve mechanical regulatory mechanisms.

Footnotes

Acknowledgements

The authors would like to thank Zachary Dean for constructive suggestions and English editing.

Declaration of Conflicting Interests

The authors declared no potential conflicts of interest with respect to the research, authorship, and/or publication of this article.

Funding

The authors disclosed receipt of the following financial support for the research, authorship, and/or publication of this article: This work was supported by National Institutes of Health Director’s New Innovator Award (DP2OD007161).

References

Risau

Flamme

Vasculogenesis. Annu Rev Cell Dev Biol. 1995, 11, 73–91.

Folkman

Angiogenesis. Annu. Rev. Med. 2006, 57, 1–18.

Hendrix

M. J. C.

Seftor

E. A.

Hess

A. R.

Vasculogenic Mimicry and Tumour-Cell Plasticity: Lessons from Melanoma. Nat. Rev. Cancer 2003, 3, 411–421.

Carmeliet

Angiogenesis in Health and Disease. Nat. Med. 2003, 9, 653–660.

Levenberg

Rouwkema

Macdonald

. Engineering Vascularized Skeletal Muscle Tissue. Nat. Biotechnol. 2005, 23, 879–884.

Koike

Fukumura

Gralla

. Creation of Long-Lasting Blood Vessels. Nature 2004, 428, 138–139.

Turing

A. M.

The Chemical Basis of Morphogenesis. Phil. Trans. R. Soc. London B Biol. Sci. 1952, 237, 37–72.

Kondo

Miura

Reaction-Diffusion Model as a Framework for Understanding Biological Pattern Formation. Science 2010, 329, 1616–1620.

Chen

T. H.

Hsu

J. J.

Zhao

. Left-Right Symmetry Breaking in Tissue Morphogenesis via Cytoskeletal Mechanics. Circ. Res. 2012, 110, 551–559.

10.

Chen

T. H.

Zhu

Pan

. Directing Tissue Morphogenesis via Self-Assembly of Vascular Mesenchymal Cells. Biomaterials 2012, 33, 9019–9026.

11.

Garfinkel

Tintut

Petrasek

. Pattern Formation by Vascular Mesenchymal Cells. Pro. Natl. Acad. Sci. U.S.A. 2004, 101, 9247–9250.

12.

Howard

Grill

S. W.

Bois

J. S.

Turing’s Next Steps: The Mechanochemical Basis of Morphogenesis. Nat. Rev. Mol. Cell Biol. 2011, 12, 392–398.

13.

Sun

Jamilpour

Wang

F. Y.

. Geometric Control of Capillary Architecture via Cell-Matrix Mechanical Interactions. Biomaterials 2014, 35, 3273–3280.

14.

Rivron

N. C.

Vrij

E. J.

Rouwkema

. Tissue Deformation Spatially Modulates VEGF Signaling and Angiogenesis. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 6886–6891.

15.

Mammoto

Connor

K. M.

Mammoto

. A Mechanosensitive Transcriptional Mechanism That Controls Angiogenesis. Nature 2009, 457, 1103–1108.

16.

Kniazeva

Putnam

A. J.

Endothelial Cell Traction and ECM Density Influence Both Capillary Morphogenesis and Maintenance in 3-D. Am. J. Physiol. Cell Physiol. 2009, 297, C179–C187.

17.

Schoen

Pruitt

B. L.

Vogel

The Yin-Yang of Rigidity Sensing: How Forces and Mechanical Properties Regulate the Cellular Response to Materials. Annu. Rev. Mater. Res. 2013, 43, 589–618.

18.

Ingber

D. E.

Tensegrity: The Architectural Basis of Cellular Mechanotransduction. Annu. Rev. Physiol. 1997, 59, 575–599.

19.

Kim

D.-H.

Wong

P. K.

Park

. Microengineered Platforms for Cell Mechanobiology. Annu. Rev. Biomed. Eng. 2009, 11, 203–233.

20.

Yang

Woolley

A. T.

Integrated Multiprocess Microfluidic Systems for Automating Analysis. J. Assoc. Lab. Autom. 2010, 15, 198–209.

21.

Wang

T. H.

Wong

P. K.

Transforming Microfluidics into Laboratory Automation. J. Assoc. Lab. Autom. 2010, 15, A15–A16.

22.

Sniadecki

N. J.

Desai

R. A.

Ruiz

S. A.

. Nanotechnology for Cell-Substrate Interactions. Ann. Biomed. Eng. 2006, 34, 59–74.

23.

Huang

H. J.

Pierstorff

Osawa

. Protein-Mediated Assembly of Nanodiamond Hydrogels into a Biocompatible and Biofunctional Multilayer Nanofilm. ACS Nano. 2008, 2, 203–212.

24.

Di Carlo

A Mechanical Biomarker of Cell State in Medicine. J. Lab. Autom. 2012, 17, 32–42.

25.

Wang

Mechanical Interactions among Cytoskeletal Filaments. Hypertension 1998, 32, 162–165.

26.

Matsumura

Wolff

Petzelbauer

Endothelial Cell Tube Formation Depends on Cadherin 5 and CD31 Interactions with Filamentous Actin. J. Immunol. 1997, 158, 3408–3416.

27.

Wakatsuki

Schwab

Thompson

N. C.

. Effects of Cytochalasin D and Latrunculin B on Mechanical Properties of Cells. J. Cell Sci. 2001, 114(pt 5), 1025–1036.

28.

Udagawa

Yuan

Panigrahy

. Cytochalasin E, an Epoxide Containing Aspergillus-Derived Fungal Metabolite, Inhibits Angiogenesis and Tumor Growth. J. Pharmacol. Exp. Ther. 2000, 294, 421–427.

29.

Wong

P. K.

Tan

C. M.

Cell Relaxation after Electrodeformation: Effect of Latrunculin A on Cytoskeletal Actin. J. Biomech. 2005, 38, 529–535.

30.

Coue

Brenner

S. L.

Spector

. Inhibition of Actin Polymerization by Latrunculin A. FEBS Lett. 1987, 213, 316–318.

31.

Harris

A. R.

Peter

Bellis

. Characterizing the Mechanics of Cultured Cell Monolayers. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 16449–16454.

32.

Phng

L. K.

Stanchi

Gerhardt

Filopodia Are Dispensable for Endothelial Tip Cell Guidance. Development 2013, 140, 4031–4040.

33.

Rosenthal

Choritz

Schlott

. Effects of ML-7 and Y-27632 on Carbachol- and Endothelin-1-Induced Contraction of Bovine Trabecular Meshwork. Exp. Eye Res. 2005, 80, 837–845.

34.

Panorchan

Lee

J. S.

Kole

T. P.

. Microrheology and ROCK Signaling of Human Endothelial Cells Embedded in a 3D Matrix. Biophys. J. 2006, 91, 3499–3507.

35.

Manser

Rho Family GTPases, Springer: New York; 2006.

36.

Chen

J. X.

SundarRaj

. Alpha-Smooth Muscle Actin Expression Enhances Cell Traction Force. Cell Motil. Cytoskel. 2007, 64, 248–257.

37.

Fenteany

Zhu

Small-Molecule Inhibitors of Actin Dynamics and Cell Motility. Curr. Top. Med. Chem. 2003, 3, 593–616.

38.

Even-Ram

Doyle

A. D.

Conti

M. A.

. Myosin IIA Regulates Cell Motility and Actomyosin Microtubule Crosstalk. Nat. Cell Biol. 2007, 9, 299–309.

39.

Mabeta

Pepper

M. S.

A Comparative Study on the Anti-Angiogenic Effects of DNA-Damaging and Cytoskeletal-Disrupting Agents. Angiogenesis 2009, 12, 81–90.

40.

Tagawa

Wang

Narishige

. Cytoskeletal Mechanics in Pressure-Overload Cardiac Hypertrophy. Circ. Res. 1997, 80, 281–289.

41.

Lau

D. H.

Xue

Young

L. J.

. Paclitaxel (Taxol): An Inhibitor of Angiogenesis in a Highly Vascularized Transgenic Breast Cancer. Cancer Biother. Radiopharm. 1999, 14, 31–36.

42.

Wang

Lou

Lesniewski

. Paclitaxel at Ultra Low Concentrations Inhibits Angiogenesis Without Affecting Cellular Microtubule Assembly. Anticancer Drugs 2003, 14, 13–19.

43.

Bagni

M. A.

Cecchi

Colomo

. Effects of 2,3-Butanedione Monoxime on the Crossbridge Kinetics in Frog Single Muscle Fibres. J. Muscle Res. Cell Motil. 1992, 13, 516–522.

44.

Sun

Foskett

. Cardiomyocyte Contractile Status Is Associated with Differences in Fibronectin and Integrin Interactions. Am. J., Physiol. Heart Circ. Physiol. 2010, 298, H2071–H2081.

45.

S. S.

Laudadio

R. E.

Lai

. Stiffness Changes in Cultured Airway Smooth Muscle Cells. Am. J. Physiol. Cell Physiol. 2002, 283, C792–C801.

46.

Fraley

T. S.

Pereira

C. B.

Tran

T. C.

. Phosphoinositide Binding Regulates Alpha-Actinin Dynamics: Mechanism for Modulating Cytoskeletal Remodeling. J. Biol. Chem. 2005, 280, 15479–15482.

47.

Ladoux

Anon

Lambert

. Strength Dependence of Cadherin-Mediated Adhesions. Biophys. J. 2010, 98, 534–542.

48.

Kniazeva

Putnam

A. J.

Endothelial Cell Traction and ECM Density Influence Both Capillary Morphogenesis and Maintenance in 3-D. Am. J. Physiol. Cell Physiol. 2009, 297, C179–C187.

49.

Olety

Wälte

Honnert

. Myosin 1G (Myo1G) Is a Haematopoietic Specific Myosin That Localises to the Plasma Membrane and Regulates Cell Elasticity. FEBS Lett. 2010, 584, 493–499.

50.

Tang

D. D.

Intermediate Filaments in Smooth Muscle. Am. J. Physiol. Cell Physiol. 2008, 294, C869–C878.

51.

Wang

R. P.

Q. F.

Anfinogenova

. Dissociation of Crk-Associated Substrate from the Vimentin Network Is Regulated by p21-Activated Kinase on ACh Activation of Airway Smooth Muscle. Am. J. Physiol. Lung Cell Mol. Physiol. 2007, 292 (1), L240-L248.

52.

Wang

R. P.

Tang

D. D.

Role of Vimentin in Smooth Muscle Force Development. Am. J. Physiol. Cell Physiol. 2006, 291, C483–C489.

53.

Mierke

C. T.

The Role of Vinculin in the Regulation of the Mechanical Properties of Cells. Cell. Biochem. Biophys. 2009, 53, 115–126.

54.

Plotnikov

S. V.

Pasapera

A. M.

Sabass

. Force Fluctuations within Focal Adhesions Mediate ECM-Rigidity Sensing to Guide Directed Cell Migration. Cell 2012, 151, 1513–1527.

55.

Thomas

W. A.

Boscher

Chu

Y. S.

. Alpha-Catenin and Vinculin Cooperate to Promote High E-Cadherin-Based Adhesion Strength. J. Biol. Chem. 2013, 288, 4957–4969.

56.

Iwanicki

M. P.

Davidowitz

R. A.

M. R.

. Ovarian Cancer Spheroids Use Myosin-Generated Force to Clear the Mesothelium. Cancer Discov. 2011, 1, 144–157.

57.

Roca-Cusachs

del Rio

Puklin-Faucher

. Integrin-Dependent Force Transmission to the Extracellular Matrix by Alpha-Actinin Triggers Adhesion Maturation. Proc. Natl. Acad. Sci. U.S.A. 2013, 110, E1361–E1370.

58.

Helfman

D. M.

Levy

E. T.

Berthier

. Caldesmon Inhibits Nonmuscle Cell Contractility and Interferes with the Formation of Focal Adhesions. Mol. Biol. Cell 1999, 10, 3097–3112.

59.

Rape

Guo

W. H.

Wang

Y. L.

Microtubule Depolymerization Induces Traction Force Increase Through Two Distinct Pathways. J. Cell Sci. 2011, 124, 4233–4240.

60.

Sarin

Gaffin

R. D.

Meininger

G. A.

. Arginine-Glycine-Aspartic Acid (RGD)-Containing Peptides Inhibit the Force Production of Mouse Papillary Muscle Bundles via Alpha(5)beta(1) Integrin. J. Physiol. Lond. 2005, 564, 603–617.

61.

Ogita

Kunimoto

Kamioka

. EphA4-Mediated Rho Activation via Vsm-RhoGEF Expressed Specifically in Vascular Smooth Muscle Cells. Circ. Res. 2003, 93, 23–31.

62.

Dai

Sodhi

Cetin

. Extracellular High Mobility Group Box-1 (HMGB1) Inhibits Enterocyte Migration via Activation of Toll-like Receptor-4 and Increased Cell-Matrix Adhesiveness. J. Biol. Chem. 2010, 285, 4995–5002.

63.

Alesutan

Seifert

Pakladok

. Chorein Sensitivity of Actin Polymerization, Cell Shape and Mechanical Stiffness of Vascular Endothelial Cells. Cell. Physiol. Biochem. 2013, 32, 728–742.

64.

Huang

Sun

. Modulation of Microvascular Smooth Muscle Adhesion and Mechanotransduction by Integrin-Linked Kinase. Microcirculation 2010, 17, 113–127.

65.

Schmitz

Nedele

Guske

. Soluble Adenylyl Cyclase in Vascular Endothelium: Gene Expression Control of Epithelial Sodium Channel-alpha, Na+/K+-ATPase-alpha/beta, and Mineralocorticoid Receptor. Hypertension 2014, 63, 753–761.

66.

Ghajar

C. M.

Blevins

K. S.

Hughes

C. C.

. Mesenchymal Stem Cells Enhance Angiogenesis in Mechanically Viable Prevascularized Tissues via Early Matrix Metalloproteinase Upregulation. Tissue Eng. 2006, 12, 2875–2888.

67.

Ghajar

C. M.

Chen

Harris

J. W.

. The Effect of Matrix Density on the Regulation of 3-D Capillary Morphogenesis. Biophys. J. 2008, 94, 1930–1941.

68.

Sieminski

A. L.

Was

A. S.

Kim

. The Stiffness of Three-Dimensional Ionic Self-Assembling Peptide Gels Affects the Extent of Capillary-Like Network Formation. Cell Biochem. Biophys. 2007, 49, 73–83.

69.

Mason

B. N.

Starchenko

Williams

R. M.

. Tuning Three-Dimensional Collagen Matrix Stiffness Independently of Collagen Concentration Modulates Endothelial Cell Behavior. Acta Biomater. 2013, 9, 4635–4644.

70.

Califano

J. P.

Reinhart-King

C. A.

A Balance of Substrate Mechanics and Matrix Chemistry Regulates Endothelial Cell Network Assembly. Cell. Mol. Bioeng. 2008, 1, 122–132.

71.

Fischer

R. S.

Gardel

. Local Cortical Tension by Myosin II Guides 3D Endothelial Cell Branching. Curr. Biol. 2009, 19, 260–265.

72.

Turturro

M. V.

Christenson

M. C.

Larson

J. C.

. MMP-Sensitive PEG Diacrylate Hydrogels with Spatial Variations in Matrix Properties Stimulate Directional Vascular Sprout Formation. PLoS One 2013, 8, e58897.

73.

Francis-Sedlak

M. E.

Moya

M. L.

Huang

J. J.

. Collagen glycation Alters Neovascularization In Vitro and In Vivo. Microvasc. Res. 2010, 80, 3–9.

74.

Rimann

Angres

Patocchi-Tenzer

. Automation of 3D Cell Culture Using Chemically Defined Hydrogels. J. Lab. Autom. 2014, 19, 191–197.

75.

Tibbitt

M. W.

Anseth

K. S.

Hydrogels as Extracellular Matrix Mimics for 3D Cell Culture. Biotechnol. Bioeng. 2009, 103, 655–663.

76.

Tse

J. R.

Engler

A. J.

Preparation of Hydrogel Substrates with Tunable Mechanical Properties. Curr. Protoc. Cell Biol. 2010, Chapter 10, Unit 10 16.

77.

Saunders

R. L.

Hammer

D. A.

Assembly of Human Umbilical Vein Endothelial Cells on Compliant Hydrogels. Cell. Mol. Bioeng. 2010, 3, 60–67.

78.

Zhao

Boudou

Wang

W. G.

. Decoupling Cell and Matrix Mechanics in Engineered Microtissues Using Magnetically Actuated Microcantilevers. Adv. Mater. 2013, 25, 1699–1705.

79.

Wang

Y. K.

Yang

M. T.

. Mechanical Regulation of Cell Function with Geometrically Modulated Elastomeric Substrates. Nat. Methods 2010, 7, 733–736.

80.

Saez

Buguin

Silberzan

. Is the Mechanical Activity of Epithelial Cells Controlled by Deformations or Forces? Biophys. J. 2005, 89, L52–L54.

81.

Dickinson

L. E.

Rand

D. R.

Tsao

. Endothelial Cell Responses to Micropillar Substrates of Varying Dimensions and Stiffness. J. Biomed. Mater. Res. A 2012, 100, 1457–1466.

82.

Kloxin

A. M.

Kasko

A. M.

Salinas

C. N.

. Photodegradable Hydrogels for Dynamic Tuning of Physical and Chemical Properties. Science 2009, 324, 59–63.

83.

Guvendiren

Burdick

J. A.

Stiffening Hydrogels to Probe Short- and Long-Term Cellular Responses to Dynamic Mechanics. Nat. Commun. 2012, 3, 792.

84.

Sunyer

Jin

A. J.

Nossal

. Fabrication of Hydrogels with Steep Stiffness Gradients for Studying Cell Mechanical Response. PLoS One 2012, 7, e46107.

85.

Chen

P. C. Y.

Herath

S. C. B.

Wang

D.-A.

. Active Manipulation of ECM Stiffness and Its Effect on Endothelial Cell Migration during Angiogenesis. Proc. World Congress Eng. 2012, I, 622–626.

86.

Junkin

Wong

P. K.

Probing Cell Migration in Confined Enivironments by Plasma Lithography. Biomaterials 2011, 32, 1848–1855.

87.

Junkin

Watson

Vande Geest

J. P.

. Template-Guided Self-Assembly of Colloidal Quantum Dots Using Plasma Lithography. Adv. Mater. 2009, 21, 1247–1251.

88.

Moon

J. J.

Hahn

M. S.

Kim

. Micropatterning of Poly(Ethylene Glycol) Diacrylate Hydrogels with Biomolecules to Regulate and Guide Endothelial Morphogenesis. Tissue Eng. Part A 2009, 15, 579–585.

89.

Raghavan

Nelson

C. M.

Baranski

J. D.

. Geometrically Controlled Endothelial Tubulogenesis in Micropatterned Gels. Tissue Eng. Part A 2010, 16, 2255–2263.

90.

Lei

Zouani

O. F.

Rémy

. Geometrical Microfeature Cues for Directing Tubulogenesis of Endothelial Cells. PLoS One 2012, 7, e41163.

91.

Uttayarat

Toworfe

G. K.

Dietrich

. Topographic Guidance of Endothelial Cells on Silicone Surfaces with Micro- to Nanogrooves: Orientation of Actin Filaments and Focal Adhesions. J. Biomed. Mater. Res. A 2005, 75A, 668–680.

92.

Myers

K. A.

Applegate

K. T.

Danuser

. Distinct ECM Mechanosensing Pathways Regulate Microtubule Dynamics to Control Endothelial Cell Branching Morphogenesis. J. Cell. Biol. 2011, 192, 321–334.

93.

Liliensiek

S. J.

Wood

J. A.

Yong

. Modulation of Human Vascular Endothelial Cell Behaviors by Nanotopographic Cues. Biomaterials 2010, 31, 5418–5426.

94.

Bettinger

C. J.

Zhang

Gerecht

. Enhancement of In Vitro Capillary Tube Formation by Substrate Nanotopography. Adv. Mater. 2008, 20, 99–103.

95.

Ingber

D. E.

Mechanical Control of Tissue Morphogenesis during Embryological Development. Int. J. Dev. Biol. 2006, 50, 255–266.

96.

Matsugaki

Fujiwara

Nakano

Continuous Cyclic Stretch Induces Osteoblast Alignment and Formation of Anisotropic Collagen Fiber Matrix. Acta Biomater. 2013, 9, 7227–7235.

97.

Milkiewicz

Doyle

J. L.

Fudalewski

. HIF-1alpha and HIF-2alpha Play a Central Role in Stretch-Induced but Not Shear-Stress-Induced Angiogenesis in Rat Skeletal Muscle. J. Physiol. 2007, 583(pt 2), 753–766.

98.

Munevar

Wang

Y. L.

Dembo

Regulation of Mechanical Interactions between Fibroblasts and the Substratum by Stretch-Activated Ca2+ Entry. J. Cell. Sci. 2004, 117(pt 1), 85–92.

99.

Riahi

Yang

Zhang

D. D.

. Advances in Wound-Healing Assays for Probing Collective Cell Migration. J. Lab. Autom. 2012, 17, 59–65.

100.

Riahi

Long

Yang

. Single Cell Gene Expression Analysis in Injury-Induced Collective Cell Migration. Integr. Bio. (Camb). 2014, 6, 192–202.

101.

Anon

Serra-Picamal

Hersen

. Cell Crawling Mediates Collective Cell Migration to Close Undamaged Epithelial Gaps. Proc. Natl. Acad. Sci. U.S.A. 2012, 109, 10891–10896.

102.

Junkin

Long

. Mechanically Induced Intercellular Calcium Communication in Confined Endothelial Structures. Biomaterials 2013, 34, 2049–2056.

103.

Long

Junkin

Wong

P. K.

. Calcium Wave Propagation in Networks of Endothelial Cells: Model-Based Theoretical and Experimental Study. PLoS Comput. Biol. 2012, 8, e1002847.

104.

Nishitani

W. S.

Saif

T. A.

Wang

Calcium Signaling in Live Cells on Elastic Gels under Mechanical Vibration at Subcellular Levels. PLoS One 2011, 6, e26181.

105.

Milkiewicz

Uchida

Gee

. Shear Stress-Induced Ets-1 Modulates Protease Inhibitor Expression in Microvascular Endothelial Cells. J. Cell. Physiol. 2008, 217, 502–510.

106.

Van Gieson

E. J.

Skalak

T. C.

Chronic Vasodilation Induces Matrix Metalloproteinase 9 (MMP-9) Expression during Microvascular Remodeling in Rat Skeletal Muscle. Microcirculation 2001, 8, 25–31.

107.

Petersen

Varoga

Zantop

. Cyclic Strain Influences the Expression of the Vascular Endothelial Growth Factor (VEGF) and the Hypoxia Inducible Factor 1 Alpha (HIF-1 Alpha) in Tendon Fibroblasts. J. Orthop. Res. 2004, 22, 847–853.

108.

Magidson

Loncarek

Hergert

. Laser Microsurgery in the GFP Era: A Cell Biologist’s Perspective. Methods Cell Biol. 2007, 82, 239–266.

109.

Reffay

Parrini

M. C.

Cochet-Escartin

. Interplay of RhoA and Mechanical Forces in Collective Cell Migration Driven by Leader Cells. Nat. Cell Biol. 2014, 16, 217–223.

110.

Supatto

Débarre

Moulia

. In Vivo Modulation of Morphogenetic Movements in Drosophila Embryos with Femtosecond Laser Pulses. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 1047–1052.

111.

Tseng

Judy

J. W.

Di Carlo

Magnetic Nanoparticle-Mediated Massively Parallel Mechanical Modulation of Single-Cell Behavior. Nat. Methods 2012, 9, 1113–1119.

112.

Shen

Tong

Bao

. Structural Responses of Cells to Intracellular Magnetic Force Induced by Superparamagnetic Iron Oxide Nanoparticles. Phys. Chem. Chem. Phys. 2014, 16, 1914–1920.

113.

Huang

Dong

C. Y.

Kwon

H. S.

. Three-Dimensional Cellular Deformation Analysis with a Two-Photon Magnetic Manipulator Workstation. Biophys. J. 2002, 82, 2211–2223.

114.

Desprat

Supatto

Pouille

P. A.

. Tissue Deformation Modulates Twist Expression to Determine Anterior Midgut Differentiation in Drosophila Embryos. Dev. Cell 2008, 15, 470–477.

115.

Sniadecki

N. J.

Anguelouch

Yang

M. T.

. Magnetic Microposts as an Approach to Apply Forces to Living Cells. Proc. Natl. Acad. Sci. 2007, 104, 14553–14558.

116.

Hsiai

T. K.

Cho

S. K.

Wong

P. K.

. Micro Sensors: Linking Real-Time Oscillatory Shear Stress with Vascular Inflammatory Responses. Ann. Biomed. Eng. 2004, 32, 189–201.

117.

Song

J. W.

Munn

L. L.

Fluid Forces Control Endothelial Sprouting. Proc. Natl. Acad. Sci. U.S.A. 2011, 108, 15342–15347.

118.

Ichioka

Shibata

Kosaki

. Effects of Shear Stress on Wound-Healing Angiogenesis in the Rabbit Ear Chamber. J. Surg. Res. 1997, 72, 29–35.

119.

Mowbray

A. L.

Kang

D. H.

Rhee

S. G.

. Laminar Shear Stress Up-Regulates Peroxiredoxins (PRX) in Endothelial Cells. J. Biol. Chem. 2008, 283, 1622–1627

120.

Ueda

Koga

Ikeda

. Effect of Shear Stress on Microvessel Network Formation of Endothelial Cells with In Vitro Three-Dimensional Model. Am. J. Physiol. Heart Circ. Physiol. 2004, 287, H994–H1002.

121.

Wang

Botvinick

E. L.

Zhao

. Visualizing the Mechanical Activation of Src. Nature 2005, 434, 1040–1045.

122.

Honarmandi

Lee

Lang

M. J.

. A Microfluidic System with Optical Laser Tweezers to Study Mechanotransduction and Focal Adhesion Recruitment. Lab Chip 2011, 11, 684–694.

123.

Conway

D. E.

Breckenridge

M. T.

Hinde

. Fluid Shear Stress on Endothelial Cells Modulates Mechanical Tension across VE-Cadherin and PECAM-1. Curr. Biol. 2013, 23, 1024–1030.

124.

Gee

Milkiewicz

Haas

T. L.

; p38 MAPK Activity Is Stimulated by Vascular Endothelial Growth Factor Receptor 2 Activation and Is Essential for Shear Stress-Induced Angiogenesis. J. Cell. Physiol. 2010, 222, 120–126.

125.

C. P.

Helm

C. L.

Swartz

M. A.

Interstitial Flow Differentially Stimulates Blood and Lymphatic Endothelial Cell Morphogenesis In Vitro. Microvasc. Res. 2004, 68, 258–264.

126.

Junkin

Leung

S. L.

Whitman

. Cellular Self-Organization by Autocatalytic Alignment Feedback. J. Cell. Sci. 2011, 124, 4213–4220.

127.

van der Meer

A. D.

Vermeul

Poot

A. A.

. A Microfluidic Wound-Healing Assay for Quantifying Endothelial Cell Migration. Am. J. Physiol. Heart Circ. Physiol. 2010, 298, H719–H725.

128.

Riahi

Dean

T. H.

. Detection of mRNA in Living Cells by Double-Stranded Locked Nucleic Acid Probes. Analyst 2013, 138, 4777–4785.

129.

Wang

J. H.

Application of Cell Traction Force Microscopy for Cell Biology Research. Methods Mol. Biol. 2009, 586, 301–313.

130.

Tan

J. L.

Tien

Pirone

D. M.

. Cells Lying on a Bed of Microneedles: An Approach to Isolate Mechanical Force. Proc Natl. Acad. Sci. U.S.A. 2003, 100, 1484–1489.

Advances in Techniques for Probing Mechanoregulation of Tissue Morphogenesis

Abstract

Keywords

Introduction

Biochemical Approaches for Mechanical Perturbation

Actin-Targeting Reagents

Microtubule-Targeting Molecules

Drugs Targeting Actin-Myosin Interactions

Targeting Intermediate Filaments

Targeting Focal Adhesions and Cell Junctions

Targeting Signaling Pathways That Regulate Cell Mechanics

Biomaterial Approaches for Modulating the Microenvironment

ECM and Substrate Properties

Geometric Control

Biophysical Approaches for Probing Tissue Morphogenesis

Mechanical Perturbation

Optical Techniques

Magnetic Technique

Shear Stress

Discussion

Footnotes

Acknowledgements

Declaration of Conflicting Interests

Funding

References