Mechano-Biochemical Regulation of Extracellular Matrix

Perspective Cite This: Biochemistry XXXX, XXX, XXX−XXX

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On Force and Form: Mechano-Biochemical Regulation of Extracellular Matrix Gwendolyn A. Hoffmann, Joyce Y. Wong, and Michael L. Smith*

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Department of Biomedical Engineering, Boston University, 44 Cummington Mall, Boston, Massachusetts 02215, United States ABSTRACT: The extracellular matrix is well-known for its structural role in supporting cells and tissues, and its important biochemical role in providing signals to cells has increasingly become apparent. These structural and biochemical roles are closely coupled through mechanical forces: the biochemistry of the extracellular matrix determines its mechanical properties, mechanical forces control release or display of biochemical signals from the extracellular matrix, and the mechanical properties of the matrix in turn influence the mechanical set point at which signals are sent. In this Perspective, we explain how the extracellular matrix is regulated by strain and mechanical forces. We show the impact of biochemistry and mechanical forces on in vivo assembly of extracellular matrix and illustrate how matrix can be generated in vitro using a variety of methods. We cover how the matrix can be characterized in terms of mechanics, composition, and conformation to determine its properties and to predict interactions. Finally, we explore how extracellular matrix remodeling, ligand binding, and hemostasis are regulated by mechanical forces. These recently discovered mechano-biochemical interactions have important functions in wound healing and disease progression. It is likely that mechanically altered extracellular matrix interactions are a commonly recurring theme, but due to limited tools to generate extracellular matrix fibers in vitro and lack of high-throughput methods to detect these interactions, it is hypothesized that many of these interactions have yet to be discovered.

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INTRODUCTION The extracellular matrix (ECM) provides cells and tissues with mechanical, structural, and biochemical signals that help direct development, guide wound healing, and maintain homeostasis. These processes are largely driven by fibrous proteins including collagens, fibronectin (Fn), elastin, tenascin, and laminin,1 and by proteoglycans, including glycosaminoglycans (GAGs) and small leucine rich proteoglycans (SLRPs).1 For example, in Xenopus development, fibronectin’s binding interactions with platelet-derived growth factor are key to guiding migrating cells.2 During hemostasis and wound healing, interactions between underlying collagen-rich ECM and platelets in blood trigger the clotting cascade leading to formation of a provisional matrix made up of fibrin and fibronectin. Then granulation tissue of fibronectin and collagen is laid down, and finally deposition of collagen-rich ECM completes the healing process.3 ECM deposition and assembly involves binding interactions between many ECM proteins, with some proteins acting as a scaffold for deposition of other proteins. For example, fibronectin guides localization of collagen fibers through their binding interactions.4 These examples show that the ECM plays a much more important role than simple structural support, and understanding biochemical interactions between the ECM and its binding partners is crucial to understanding key processes in health and disease. It has become apparent that the structural and biochemical roles of the ECM are linked; force-mediated biochemical interactions play a significant role in these processes. However, only fibronectin force-dependent binding has been extensively © XXXX American Chemical Society

investigated, mostly due to a lack of methods to study other proteinaceous fibers under force. Even so, only a few binding partners of fibronectin have been evaluated for force regulation due to lack of high-throughput methods to identify these interactions, and most force-mediated biochemical interactions discovered in vitro have not yet been demonstrated in vivo. The matrix as a whole can be deformed under force, and so can the individual proteins that make up the matrix. Thus, ample motivation exists to characterize the mechanical features of ECM. Most ECMs are mechanically loaded like a spring, and in some tissues, such as bone,5 such loading is a necessary signal for normal tissue development and maintenance. Moreover, the ECM acts as a reservoir for growth factors like vascular endothelial growth factor A (VEGF-A165) and transforming growth factor beta 1 (TGF-β1), which can aid in tissue repair and wound healing: mechanical properties of the matrix can determine when these growth factors are released from the matrix.6−9 ECM mechanics are consequently an important regulator of cell signaling and function, and these mechanics are in turn influenced by the process of ECM assembly. To characterize the properties of ECM structures, one must either measure natural ECM produced by cells or, alternatively, reconstruct simplified ECM structures for in vitro interrogation. Special Issue: Mechanical Forces in Biochemistry Received: March 14, 2019 Revised: May 29, 2019 Published: May 30, 2019 A

DOI: 10.1021/acs.biochem.9b00219 Biochemistry XXXX, XXX, XXX−XXX

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Figure 1. Examples of ECM assembly requiring coordination of multiple binding partners. (A) Fn in its compact conformation binds to integrins, which exert force on the Fn molecule. (B) Fn is stretched into its extended conformation, and (C) possibly further stretched to unfold a domain. This conformation change allows binding of other Fn molecules and self-assembly of Fn matrix.14 (D) In the endoplasmic reticulum, some of the collagen peptide’s proline residues are hydroxylated to form hydroxyproline (see inset). (E) Three peptide chains combine in a helix to form procollagen. (F) The procollagen is packaged for export at the Golgi apparatus. (G) Procollagen is converted to collagen when enzymes cleave the procollagen peptides (see inset) at the N- and C-termini.15 (H) Collagen microfibrils assemble with the aid of collagen V, forming a banded pattern.16 (I) SLRPs bind to the microfibrils and (J) encourage collagen fibrils to grow in length before they (K) grow laterally.17 (L) Collagen fibrils bind to relaxed fibronectin fibers, which guide matrix assembly.4

Synthetic ECM fibers can be generated in vitro using several methods that vary in complexity and throughput, but most methods are limited in the type of ECM protein that can be used; therefore, it is not surprising that some ECM proteins have not yet been incorporated into synthetic fibers. Mechanical characterization of synthetic fibers is especially useful due to challenges associated with extracting and working with natural ECM fibers in vivo or with cell culture systems. The importance of cell substrate elasticity or modulus (a measure of matrix stiffness) has been shown in numerous in vitro studies, as it controls fundamental cell properties such as migration10 and differentiation.11 A property related to matrix stiffness is extensibility. A matrix is generally either soft and extensible, or it is stiff and not extensible.12,13 The extensibility of a matrix is a measure of how far the matrix can be stretched before breakage and may be important for matrix resilience.

(e.g., resistance to compression or tension) in the same tissue, enable tissues to stiffen with increasing loads as fibers are stretched and recruited, and focus stress on some structures while protecting other structures such as cells.21,22 Frequently, cell assembly of ECM requires force application and/or additional proteins interacting with the primary ECM type. For example, Fn matrix assembly requires force application by cells. Fn matrix is critical due to its role as a scaffolding material for subsequent deposition of other matrix components. Cells bind to fibronectin via integrins and apply force to Fn through actin cytoskeleton contraction, stimulated by Ras homolog (Rho). This unfolds a region of Fn to expose a cryptic Fn binding domain that enables matrix self-assembly (Figure 1A− C).14 Type I collagen assembly begins inside the cell. At the endoplasmic reticulum, some lysine and proline residues are hydroxylated, and three α-chains intertwine into a right-handed helix, forming procollagen (Figure 1D,E). The procollagen is prepared for export in the Golgi apparatus, then moved to the fibripositors, which are inward folds/protrusions of the cell membrane that aid in collagen deposition during development.23 Fibripositors, which are stabilized by actin and require cell tension to form, are thought to aid in collagen fibril alignment in tendon.24,25 Between the Golgi apparatus and deposition, procollagen’s N and/or C terminal propeptides are enzymatically cleaved resulting in collagen (Figure 1F,G).15 Enzymes including bone morphogenic protein-1, a disintegrin and metalloproteinase with thrombospondin motifs (ADAMTS), mephrin α and β, or tolloid-like proteases, cleave the collagen propeptides. Lack of propeptide cleavage impairs collagen assembly into fibrils, leads to irregular collagen organization, smaller and more loosely packed fibrils, and

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GENERATION OF ECM IN VIVO AND IN VITRO Assembly of ECM in Vivo Requires Coordination of Multiple Players. To study the ECM, methods to obtain or generate physiologically relevant ECM are necessary. One source is cell-assembled ECM (Figure 1), obtained from cells in vitro or tissues in vivo. In vivo, ECM assembly involves binding between molecules to form interconnected structures that vary between tissues. Much of the ECM is fibrous, which is important, since the presence of fibers allows transmission of forces over long distances, enabling cells to sense each other and communicate mechanically over distances of up to 100 μm.18,19 It also allows for independent local and global remodeling of the matrix in response to different stimuli.20 Fibrous matrix is important mechanically, because a composite network of fibrils can provide different mechanical functions B


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linking activity.31 Cross-linking and other ECM modifications have a strong impact on ECM properties. After ECM is generated, it can undergo a variety of posttranslational enzymatic modifications that affect its assembly, function, and mechanical properties. One such post-translational modification is citrullination, in which a peptidylarginine deiminase (PAD) converts arginine to citrulline. Since citrulline is less basic and less positively charged than arginine, this change can alter protein folding and interactions. Citrullination of proteins can affect many proteins in disease including myelin basic protein (associated with multiple sclerosis)32 and vimentin (associated with liver fibrosis);33 some ECM proteins it impacts are fibrin, fibronectin, and collagen. For example, fibrin citrullination reduces the ability of plasmin to break down fibrin plaques, which likely contributes to the progression of rheumatoid arthritis (RA).32 Enzymes that work primarily to mediate collagen modifications are lysyl hydroxylase (LH), which converts collagen lysine residues into hydroxylysine, and lysyl oxidase (LOX), which converts collagen and elastin lysine residues to allysine and hydroxylysine residues to hydroxyallysine to generate reactive aldehyde groups. Allysine can react with hydroxylysine, hydroxylallysine with hydroxyallysine, and hydroxylysine with lysine to form divalent cross-links, which can react with each other to form mature pyrrole and pyridinoline trivalent cross-links. The cross-linking reactions that occur vary by tissue type, but they are essential for fiber stability and appropriate extracellular matrix stiffness.34−36 LOX also contributes to development of tissue fibrosis, which can increase tumor metastasis in fibrosis-affected tissues.35 LH is another contributor to fibrosis. A specific type of LH, LH2B, hydroxylates lysine residues in collagen’s telopeptides. The hydroxylated lysine reacts with lysine, hydroxylysine, or histidine in collagen to form a pyridinoline cross-link, which is increased in fibrosis. Interestingly, the authors that found an increase in LH2B did not show an increase in LOX in fibrosis, which suggests that multiple, separate mechanisms could lead to fibrosis.37 Tissue transglutaminase is another important enzyme, which cross-links glutamine residues with primary amines (such as the primary amine on lysine residues) to form stable isopeptide bonds. Tissue transglutaminase is known to act on many ECM proteins including collagen, fibronectin, fibrinogen, and laminin, where it is thought to stabilize and help prevent degradation of these proteins. In disease, type 2 tissue transglutaminase has been shown to play a role in liver cirrhosis, liver, renal, and lung fibrosis, rheumatoid arthritis, and osteoarthritis.38,39 Independent of its catalytic activity, tissue transglutaminase type 2 forms a complex with fibronectin, which facilitates cell adhesion independent of RGD (arginineglycine-aspartic acid) binding and enhances fibronectin deposition.40 Binding interactions between proteins can also play a role in modulating ECM mechanics similar to crosslinkers by coupling multiple ECM proteins. For example, type XII collagen contributes to the mechanical properties of the ECM by binding to tenascin-X, decorin, fibromodulin, and cartilage oligomeric matrix protein/thrombospondin-5. As tenascin-X and decorin bind to collagen fibrils, collagen XII binding forms a bridge between collagen fibrils. Collagen XII, upregulated in fibrosis and cancer, is thought to be regulated by TGF-β1 and/or tensile strain.41 In a functional assessment of tendon during chick development, collagen cross-linking brought about by LOX was shown to be the primary contributor to tendon mechanical properties, rather than collagen microstructure, or collagen, cell, or GAG content.42

lower skin rupture force. If the N-terminal propeptide cannot be cleaved in vivo, this results in Ehlers-Danlos syndrome (EDS) type VII.26 After propeptide removal, collagen triple helices assemble into microfibrils at the cell surface (Figure 1H).15 Collagen V plays a critical role in the initiation of fibril assembly and control of fibril diameter, likely due to its shape. Without collagen V, collagen I forms abnormal, large, and irregular aggregates, leading to Classical EDS in patients with collagen V mutations.16 After microfibril assembly and deposition, the microfibrils assemble into larger fibrils with assistance from other ECM molecules. Collagen interactions with SLRPs and Fn help organize its ECM structure. SLRPs including lumican, fibromodulin, and biglycan bind to collagen fibrils and aid in assembly of larger fibrils from smaller fibrils. Initially, SLRPs lumican and fibromodulin prevent lateral fibril growth, so that fibrils can initially grow longer (Figure 1I,J). Later, SLRPs encourage lateral growth and fusion of fibrils (Figure 1K). SLRP deficiency can lead to decreased tendon stiffness, fragile skin, and loose joints.17 Lack of lumican leads to corneal opacity and an eightfold decrease in skin tensile strength due to formation of disorganized, loosely packed collagen fibrils with large diameters.27 Attached to SLRPs are charged GAGs, which bind water and help the ECM resist compressive forces.17 Collagen deposition is aided by Fn matrices: Collagen I colocalizes with relaxed Fn fibers, binding to Fn’s collagen and gelatin binding site. This means that the initial Fn matrix determines where collagen matrix will be deposited (Figure 1L)4 in collagen I-rich ECMs. Force also plays an important role in collagen organization; cell contractility leads to alignment of collagen fibers and the formation of collagen tracts.19 An alternative theory to fibripositor and force alignment of collagen is liquid crystal induced alignment. Collagen is thought to act like a liquid crystal due to its high aspect ratio, which in liquid crystal solutions causes molecules to align above a critical concentration, and the presence of liquid crystal-like structures found within collagen in vitro and in vivo. Yet, it has not been demonstrated that collagen’s liquid crystalline properties cause alignment in vivo.28 In other types of ECMs, such as basement membranes (BMs) briefly described below, protein shape and protein interactions regulate ECM deposition. An important and specialized ECM structure is the basement membrane, which lines tissues and acts as a barrier between different compartments. Laminin, which is essential to BM assembly, is a cross-shaped molecule with one long arm and three short arms. The shape is formed from α, β, and γ chains twisted together. Laminins bind to the cell surface with their long arm, either directly through integrins or indirectly through sulfated glycolipids, α-dystroglycan, or heparan sulfate proteoglycans. The three short arms can bind to each other and allow polymerization of a sheetlike laminin network.29 Laminin has been shown to coordinate with substrate mechanical properties to alter vascular smooth muscle cell’s response to stiffness.30 This suggests force could play a role in laminin’s function. In BM maturation, collagen IV binds and forms a network of its own to stabilize the basement membrane. Collagen IV binding to the BM likely occurs through heparan sulfate proteoglycans (perlecan and agrin) as well as nidogen, which binds laminin’s short γ-chain arm.29 Lysyl oxidase-like protein-2 (LOXL2) has been shown to be required for collagen IV network assembly in the basement membrane through an unknown mechanism that is not related to LOXL2’s crossC


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Biochemistry This shows that post-translational modifications are important to tissue properties and function. Large proteoglycans and GAGs contribute to ECM function and mechanics. Hyaluronic acid, or hyaluronan, is a nonsulfated, linear, high molecular weight GAG with viscoelastic (time-dependent) properties resulting from chain entanglement. Its functions include maintaining tissue hydration by immobilizing water, lubricating joints, providing resistance to compression, preventing angiogenesis, and maintaining tissue integrity. Lower molecular weight fragments of hyaluronan seem to indicate tissue damage and are angiogenic, inflammatory, and can stimulate fibroblasts to produce collagen.43,44 Hyaluronan also associates with large chondroitin sulfate proteoglycans such as aggrecan and versican. Aggrecan has many GAG side chains that are charged, causing it to swell with water, which helps cartilage resist compression and contributes (along with collagen II) to the viscoelasticity of cartilage.45 Similarly, versican increases matrix viscoelasticity, which supports cell proliferation and migration.46 ECM assembly in vivo is highly complex and can be modulated by many components, resulting in altered mechanical properties. Some aspects of ECM assembly have been shown to be modulated by forces; perhaps some of the interactions discussed above are also influenced by force through as of yet undiscovered mechanisms. In Vitro Tools for Generating ECM. Sources of cellassembled ECM include decellularized tissues or ECMs derived from cells cultured in vitro. The advantage of cell-assembled ECMs is that they are likely the most physiologically relevant, since they have similar ECM architecture and composition to ECMs in vivo.51,54 There are many properties of cell-derived ECM that are difficult to study due to the complexity of the ECM and the entanglement of multiple types of ECM fibers. Thus, there has long been a need to produce synthetic matrix fibers that can be interrogated in physiologically relevant systems; these synthetic matrices have a variety of uses in regenerative medicine and tissue engineering. Numerous methods of synthetically manufacturing ECM (Figure 2) have been developed. Fibronectin fibers can be generated in a simple but low-throughput method called fiber pulling. A concentrated droplet of fibronectin is placed on a surface, and a sharp object such as a needle or pipet tip is dragged out of the droplet. A 2−5 μm fiber forms as the tip moves away from the air−liquid interface.53 Likewise, a low-throughput, simple method to generate collagen fibers occurs through confinement and crowding. High-concentration collagen solutions are produced by dialyzing collagen against poly(ethylene glycol) (PEG) solutions. The concentrated collagen is confined between two glass coverslips and incubated at 37 °C to produce aligned lamellae of collagen fibers.52 Other methods are more versatile and allow customization of ECM compositions. For example, in wet spinning, a protein solution is pumped out of a syringe into a bath of alcohol, where fibers (generally in the micrometer size range) form as water diffuses out of the stream of liquid into the alcohol. Wet spinning is simple to set up, high throughput, and versatile.49 Electrospinning is similar to wet spinning, but it has a more complicated setup that yields narrower fibers in the nanometer range. In electrospinning, a protein solution is extruded out of a syringe toward a metal collecting plate. A high voltage (15−30 kV) is applied between the syringe needle and the collecting plate, which causes nanofibers to form from the stream of liquid and deposit on the collecting plate. A disadvantage of electrospinning proteins is that the protein

Figure 2. In vitro methods for generating ECM fibers. Methods for generating fibers range from complicated to simple and from low to high throughput. In microfluidic spinning, a protein solution flows through the main channel, while solutions from the crossing channels focus the flow into a fiber. The fiber exits the device into a wash bath, where it can be wound around a mandrel.47 In electrospinning, a protein solution is loaded into a syringe, and a voltage is applied between the syringe needle and a collecting plate to form fibers.12,48 In wet spinning, the protein solution is loaded into a syringe and extruded into a coagulation bath.49 Cell-assembled ECM is arranged by cells in vitro or in vivo and harvested by removing the cells.50,51 In collagen crowding and confinement, a highly concentrated collagen solution is placed between coverslips, and collagen fibers form spontaneously.52 In fiber pulling, a sharp tip is dipped into a droplet of fibronectin and pulled away slowly, which results in fiber formation between the tip and the droplet surface.53

generally must be dissolved in a harsh organic solvent, such as hexafluoro-2-propanol. Nevertheless, fibrinogen and collagen fibers can be spun using electrospinning.12,48 To generate larger fibers of silk and collagen in the micron size range, a microfluidic device can be used. The device has an inlet for the protein solution to be spun and two inlets that intersect the first inlet perpendicularly to introduce a flow focusing solution. The solution contains PEG to match the viscosity of the protein solution and precipitate the fiber, and its flow rate determines the diameter of the fibers.47,55 The wide range of available methods allows choice in complexity, throughput, fiber composition, and diameter. The methods listed above are a step closer to recapitulating the in vivo ECM microenvironment compared to typical cell culture conditions. While cells are often cultured on gels or coatings of ECM proteins, these gels and coatings likely do not adequately capture the complex architecture of the ECM or the topological and mechanical signaling that it provides. Generating fibrous ECM recreates some of the complexity of in vivo ECM with its ability to transmit forces over long distances, and many existing techniques allow control of the ECM composition and fiber diameter. These techniques are advantageous, because they are able to generate large amounts of ECM with defined properties in a reproducible manner. However, lack of knowledge about the molecular architecture D


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Figure 3. Measuring and calculating ECM mechanical properties. (A) Methods used to make measurements. Ranges below technique names indicate force or stress ranges that can be measured with each technique. AFM is used to stretch a fiber laid across a groove between two ridges.12 The deformation of the fiber and the deflection of the cantilever are used to determine mechanical properties. The optical fiber/micropipette-based tester uses one optical fiber/micropipette to stretch the ECM fiber and the deflection of the other to measure the force in the ECM fiber laid between them.56 The Hall-effect based tester uses a motor to stretch the sample and voltage generated by the deflection of a beam with an attached Hall-effect sensor in a magnetic box to determine the force in the sample.57 In rheology, one part, such as a plate, rotates against the sample, and the rotational response of another plate in contact with the sample is measured to obtain mechanical properties. In tensile testers, load cells accommodating a wide range of forces are used to measure the forces in the sample, while a motor stretches the sample. (B) An example stress−strain curve shows how mechanical properties are calculated. The strength is the maximum stress attained, the modulus is the slope of the linear region, the toughness is the area under the curve, and the strain at which the fiber fails or breaks is the extensibility. (C) A comparison of stress−strain curves from two different methods of generating and measuring collagen fibers. Both measurements were made on dry fibers in air.48,55,58

fiber by moving the cantilever along the groove, perpendicular to the fiber. On the basis of the amount the fiber is stretched and the deflection of the AFM cantilever, the stress and strain in the fiber can be calculated.12,48 A method to measure the next smallest forces, down to nanonewtons, was developed in our lab. In this method, pulled micropipettes or sections of optical fiber act as actuators and sensors. One micropipette or optical fiber is attached to a motor and acts as a puller or actuator, and the other is fixed to a mount and acts a sensor. A fibronectin fiber is generated between micropipettes through pulling, or a wet spun fiber is attached to optical fibers with epoxy before submersion in fluid. The actuator stretches the fiber and is used to calculate fiber strain, while the deflection in the sensor is used to measure the force within the fiber. In the optical fiber setup, the force range measured can be easily adjusted by changing the length or diameter of optical fiber used.56 Another tensile testing method to measure slightly larger forces uses a Halleffect sensor attached to a beam. The beam’s stiffness determines the mechanical force range that can be measured, within the micronewton range. In this setup, the sample is attached between a motor actuator and the beam. The motor stretches the sample, which deflects the beam and the Halleffect sensor attached to the beam. The change in position of the Hall-effect sensor within a magnetic yoke registers a proportional change in voltage, which can be used to calculate the force in the sample. The motor data and the Hall-effect data are used to generate stress−strain curves for mechanical analysis.57 For bulk ECM structures, commercially available rheology or tensile testing equipment can be used. Magnetic resonance elastography (MRE) has recently been used to determine the viscoelastic properties of collagen gels, suggesting that it is likely suitable for mechanical testing of

and molecular conformations limits the utility of studies that use these synthetic fibers. This is an area that is ripe for future innovation and discovery, as synthetic ECM fibers are useful for many applications and studies. Possible methods of identifying molecular architecture and conformation are included in the Characterizing ECM section below. Additionally, most methods of synthetically generating ECM fibers currently only use one type of ECM protein, which does not match the compositional complexity of native ECM. Future methods should strive to incorporate more proteins to more closely recapitulate the in vivo environment and perhaps provide cells with more physiologically appropriate signals.

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CHARACTERIZING ECM Mechanical Characterization. The ability to characterize cell-generated or artificially manufactured ECM is important, as it allows comparison of mechanical properties, composition, conformation, and structure that may occur due to different manufacturing methods, physiological or pathophysiological conditions, or strain state. Mechanical property measurement (Figure 3A) generally requires means of applying a strain and measuring the force resulting from the applied strain or vice versa. To measure individual ECM fibers, various forms of tensile tests have been developed. The method chosen to test a specific sample depends on the force required to break the sample and the configuration of the sample. Starting with the method used to measure the weakest forces, atomic force microscopy (AFM) was adapted to measure the mechanical properties of soft fibers. In this method, fibers are laid over a patterned surface of grooves and ridges. In places where a fiber is suspended between two ridges over a groove, an AFM cantilever is brought into contact with and used to stretch the E


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liquid chromatography. Next, tandem mass spectrometry is performed, and software along with a database of proteins is used to identify proteins present in the original sample based on peptide fragment mass-to-charge ratio. Hundreds of proteins at a time can be identified using mass spectrometry, which allows a far more extensive characterization of ECM composition than is possible with other techniques, such as identifying ECM composition differences between tissues and showing that ECM composition changes with the metastatic potential of the tumor. This technique has also been used to identify the origin of tumor ECM in mouse xenograft models, since the species origin of proteins can be identified with species-specific peptide domains. They found that tumor ECM is derived from both the tumor and the surrounding stroma.65 The large amount of information gained from this method could make the lengthy sample preparation worthwhile, and it seems likely that this technique will continue to yield important discoveries about ECM composition. Other proteomics techniques can be used to determine tissue and ECM composition, such as matrixassisted laser desorption ionization imaging mass spectrometry (MALDI IMS), which has been used to determine the composition of tissues including quantification of collagens, elastin, and proteoglycan content and also provides protein spatial localization information.66,67 ECM composition is important, but how the proteins are organized is just as important. Structural and Organizational Characterization. Structural analysis identifies the conformation of proteins and is defined by degree of protein unfolding and matrix organization. Fourier transform infrared (FTIR) spectroscopy and circular dichroism (CD) can both be used to identify relative amounts of α-helix, β-sheet, and random conformations in proteins.68,69 Circular dichroism, as well as NMR or fluorescence, can be used with a stopped-flow setup to study kinetics of protein folding or unfolding. In this setup, a denaturant is flowed in to a native protein to evaluate protein unfolding, or a refolding buffer is flowed in to a denatured protein to evaluate protein refolding. CD, NMR, or fluorescence is used to monitor the conformation of the protein as it folds or unfolds, and the resulting curves give information about folding kinetics. This technique has been used to observe that the 10th type II module of fibronectin refolds quickly.70 To observe collagen assembly from monomers into fibrils, FTIR71 or collagen turbidity72 can be used. Collagen fibril organization can be investigated using polarized light microscopy. Collagen alignment gives the fibrils birefringent properties and produces a signal under microscopy using polarized light, so the technique allows evaluation of whether or not fibrils are aligned.73 An additional collagenspecific technique is second harmonics generation microscopy, which allows visualization of collagen fibrils and fibers without any labeling due to collagen’s hyperpolarizability property. This technique can also be used to determine collagen organization and alignment,74 and it has been combined with endogenous multiphoton microscopy signals to evaluate ECM remodeling in lung fibrosis. Another multimodal, label-free imaging technique that combines coherent anti-Stokes Raman scattering (CARS), sum-frequency generation (SFG), and two-photon excitation fluorescence (TPEF) has been used to visualize arterial ECM. CARS shows collagen and elastin, and SFG and TPEF show collagen and elastin, respectively, to validate the CARS measurements.75 The molecular density of the ECM makes microscopy difficult, so there is much potential for development of new techniques or new applications of existing

other bulk ECM structures as well. In this method, the sample deflection in response to waves of various frequencies is detected by magnetic resonance imaging and used to calculate storage and loss modulus of the sample.59 Mechanical tests from each testing method result in a stress− strain curve (Figure 3B). From this curve, important information about the material tested is gleaned. The slope of the linear region is the modulus or stiffness, the strain at failure is extensibility, the maximum stress is tensile strength, and the area under the curve is toughness. For example, collagen fibers produced using microfluidic spinning and tested using a tensile tester have a modulus of ∼4 GPa, strength approaching 400 MPa, extensibility of 25%, and toughness of ∼50 MJ m−3.55 Collagen fibers formed through electrospinning and tested using AFM have a modulus of ∼3 GPa, strength of 25 MPa, and extensibility of 33% (Figure 3C).48 Even though these fibers are both generated from collagen type I, their mechanical properties (most notably strength) are different. This may be due to structural differences due to the different methods used to produce the fibers, as well as the different testing methods used. One notable difference between these examples is the lower standard deviation in measurements for the electrospun, AFM-tested fibers, which indicates either a lower variability in spinning or less noise while testing the fibers. Both fibers were tested in a dry state, which, while helpful for comparison, will differ from tests performed on wet, hydrated fibers that more closely match fibers in vivo. This is a common issue that should be remedied in the future by new methods to test hydrated fibers. Mechanical properties of ECM can also be impacted by modifications made to them after assembly. For example, crosslinking collagen with lysyl oxidase increases modulus and strength.60 The collagen fibers above were tested with tensile tests, which means that force in the fiber is measured as the fibers are stretched until they break. As a fiber breaks, the abruptness with which stress returns to zero can be used to evaluate the mode of failure. A clean drop to zero stress indicates brittle failure, in which the material has separated all at once. A gradual drop indicates ductile failure, where the material has adsorbed energy and stretched before failure.61 Both types of collagen fibers above appear to undergo primarily brittle failure. Other types of tests might better approximate the types of forces a protein could experience in vivo, such as a constant force (creep) experiment. In this setup, a constant force is applied to the fiber, and its response over time is measured.56 Mechanical testing is important, since ECM has many structural functions, but mechanical characterization is just one of many ways to evaluate ECM. Characterization of Composition. ECM composition is important to characterize, since it can vary from tissue to tissue and between normal and disease states. Hydroxyproline assays can be used to evaluate collagen content,62 since hydroxyproline is predominantly found in collagen. Content of sulfated GAGs can be measured with a dimethylmethylene blue (DMMB) assay, in which the DMMB absorption spectrum changes when sulfated GAGs are bound.63,64 Naba and Hynes developed a proteomics technique combining ECM enrichment, mass spectrometry, and bioinformatics to characterize the matrisome, which is the collection of ECM and ECMassociated proteins present in an organism. To enrich the ECM, ECM components are separated from other contaminants in a tissue sample such as cellular proteins. Then the ECM enriched fraction is solubilized and reduced, alkylated, deglycosylated, digested, separated by isoelectric point, and finally separated by F


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predict cell spreading on viscoelastic substrates.85 Predicting and understanding protein unfolding can help lead to the identification of new ways that the ECM regulates cells and is regulated by forces. However, modeling the ECM as springs and dashpots may be too simplistic; for example, it does not capture the ability of protein domains to make a rapid transition from a folded to unfolded state after reaching a threshold force value. On the one hand, we and others have modeled fibronectin fiber mechanics as determined by chain extension and module unfolding, but these models are limited to fibers composed only of a single molecular component.81−83,86 Steered molecular dynamics, on the other hand, are limited by the amount of time it takes to run the simulations, as it can take one week to simulate one nanosecond of time.78 This also limits the number of molecules that can be incorporated into the simulation, as more molecules would require more time to simulate.

techniques in this area. One possible new application of an existing technique to the ECM is expansion microscopy. In this technique, a sample is labeled or stained with antibodies, embedding in and cross-linked to a swellable polyelectrolyte gel, digested with proteolytic enzymes, swelled in water, and imaged with a confocal microscope to obtain super-resolution images of the original sample’s labeled components.76 This seems like a promising technique for investigation of nanoscale ECM structure. Imaging techniques continue to be the subject of further development and innovation, and advancements in this field are predicted to increase our understanding of matrix functions. A specific subset of structural analysis is determination of the strain state of the matrix or matrix proteins, which measures the degree of protein unfolding due to stretching. Fibronectin has perhaps been most extensively studied due to the large extensibility of Fn fibers of up to fourfold or larger strains at failure and the finding that Fn fibers in vivo and in cell culture are also stretched to failure by cell contractile forces. Fn’s strain state can be determined several ways. One method is using Förster resonance energy transfer (FRET) sensors: fibronectin is chemically denatured, then FRET acceptors are bound to cysteines, and FRET donors are bound to lysines. The fibronectin refolds when placed in buffer. When the fibers are stretched, the FRET ratio between the fluorescence of the acceptor and the donor decreases as cysteines and lysines are separated from each other. The FRET ratio can be used to calculate the degree of strain in the fibronectin fibers.53 Phage display is another useful technique that has been used to identify peptides that are sensitive to the degree of strain in fibronectin fibers. A fluorescent molecule is attached to the peptide-displaying phage, or the peptide is conjugated to a quantum dot, and the degree to which two different probes bind fibronectin is used to calculate fiber strain. This method is advantageous, because it does not require chemical labeling of fibronectin prior to the experiment, which allowed detection of strain state in cell-assembled ECM and ex vivo lung slices.77 Methods to discriminate strain state of other protein fibers, without directly applying a specific strain to the protein fiber, are limited. Modeling can be used to identify potential protein unfolding mechanisms and predict ECM mechanics. Steered molecular dynamics (SMD) is a method that can be used to simulate force-induced unfolding of a protein. The simulation starts with known protein structures solvated in water and equilibrated to a certain temperature. One end of the protein is fixed, while a force is applied to the other end at a constant velocity or force. This allows information to be gained about the force required to unfold the protein and mechanism through which the protein likely unfolds. This method has been used to investigate the unfolding of fibronectin domains, titin’s Ig domains, and several other proteins.78,79 Other simulations, most of which model ECM structures as a combination of springs and dashpots, can be used to make predictions about the mechanics of ECM components and structures.80−83 To model viscoelastic (time-dependent) behavior, Maxwell, Kelvin−Voigt, or Burgers models can be used. The Maxwell model consists of a spring and dashpot connected in series, which tends to be more accurate under stress relaxation (constant strain) conditions, whereas the Voigt model is made up of a spring and dashpot in parallel and is more accurate under creep (constant stress) conditions.84 The combination of a Maxwell and Voigt model in series results in the Burgers model, which has been used to

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MECHANO-BIOCHEMICAL ECM REGULATION Forces Regulate ECM Remodeling. The ECM is regulated by force (Figure 4), and ECM properties in turn

Figure 4. ECM binding is regulated by strain. When fibrin fibers are relaxed, plasmin can bind and cleave fibrin. Under strain, the binding site is destroyed, and plasmin can no longer bind or cleave fibrin.87 When fibronectin is relaxed, IL-7 is unable to bind, but under strain, the second and third type I domains are stretched away from each other, allowing IL-7 to bind.7 Both α5β1 and αvβ3 integrins can bind to relaxed fibronectin. Under strain, the RGD loop on FnIII10 is separated from the synergy binding site on FnIII9, which leads to a decrease in α5β1 binding and an increase in αvβ3 binding.50

regulate cellular functions. The deformation of ECM under force plays a key role in regulating its remodeling; these forces arise from cell traction, fluid flow, and macroscopic movement such as breathing. Cells can apply forces in the nanonewton range,88 and shear stresses in arteries can reach up to 70 dyn/ cm2.89 Fibronectin fibers are known to unfold in response to applied force through multiple mechanisms, which gives Fn its high extensibility. Fn can convert between a compact conformation that is stabilized by electrostatic bonds between domains and an extended conformation with the electrostatic bonds broken. Fn type III domains can also unfold when hydrogen bonds that stabilize its beta sandwich structure are broken.90 The first type III module of fibronectin has a cryptic self-assembly domain that is exposed due to force application and aids in Fn matrix formation.14,91 Fn unfolding has been G


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Biochemistry observed through AFM,90 antibody labeling,14 FRET labeling,53 and direct labeling of cysteine residues that are normally buried in the equilibrium fold of a subset of Fn type III modules.92,93 Tenascin also contains fibronectin type III domains and unfolds when stretched with an AFM at forces of ∼137 pN.94 Several laminin domains unfold when stretched via AFM.95 It is possible that the unfolding of these ECM molecules could also have important regulatory consequences. In collagen, collagenase will preferentially degrade relaxed, unloaded fibers over mechanically loaded, strained fibers.73,96−98 This would help preserve load-bearing structures, while removing unnecessary fibers that are not load-bearing. Similarly, fibrin fiber lysis by plasmin in vitro is reduced when the fibers are under strain (200%−250%), possibly due to strain-induced alterations to the plasmin cleavage sites on fibrin. While this effect has not yet been observed in vivo, it is possible that blood flow, platelets, cells, clot retraction, or fiber prestrain could strain fibrin fibers during wound healing. Fibrin crosslinking also reduces fiber lysis, but surprisingly, to a lesser extent than fiber strain.87 It seems likely that this mechanism may also apply to other ECM fibers. One obstacle to overcome is that it is often difficult to study ECM fiber biochemistry while simultaneously stretching ECM fibers in a controlled manner. Thus, new tools that allow both measurement of ECM biochemistry and mechanics would open numerous new studies of ECM mechanobiology. Ligand Binding Is Regulated by ECM Strain. Forces and strain in the ECM also regulate the binding of ECM accessory proteins that alter their functions or catalyze their breakdown or remodeling. The N-terminal type I modules of fibronectin have strain-dependent binding to at least two ligands. On the one hand, interleukin-7 (IL-7) is a cytokine that plays a role in Tcell and lymphatic system regulation and shows increased binding to strained fibronectin fibers compared to relaxed fibers.7 On the other hand, bacterial adhesins show decreased binding to fibronectin’s N-terminal region when fibronectin is under strain due to disruption of its binding site. This mechanosensitive binding of bacterial adhesins might regulate bacterial adhesion and infection in healthy versus wounded tissue.99 Fibronectin strain also alters integrin binding. Integrins bind to the RGD loop on the 10th type III module of fibronectin; however, some integrins also bind to the peptide sequence PHSRN on the adjacent ninth type III module, also known as the synergy site. Under strain, the distance between these binding sites increases, which leads to a decrease in binding of integrins (e.g. α5β1) stabilized by the synergy site and an increase in binding of integrins that only bind RGD (e.g., αvβ3).50 Since integrins are transmembrane receptors, this integrin switching could alter cell signaling and behavior. Indeed, our lab has shown that fibronectin fiber strain reduces its ability to serve as an adhesive substrate for cell attachment and migration.100 Transforming growth factor beta 1 (TGF-β1) can be released from the ECM due to force application. TGFβ1 is secreted in a latent form, bound to the latency associated peptide (LAP) and in a complex with latent TGF-β-binding protein 1 (LTBP-1). LTBP-1 binds the complex to the ECM, and integrins can bind to the complex through RGD domains on LAP. If a cell pulls on the complex through its integrin linkage to LAP with enough force (∼40 pN), and the LTBP-1 is linked to ECM that is stiff enough, the LAP domain holding TGF-β1 will unfold and release TGF-β1 from the ECM.101 If the underlying ECM is already under stress or strain, release of TGF-β1 is increased compared to unstrained matrices.102 This

is important, because TGF-β1 signaling is a major contributor to fibrosis. ECM strain is thus a strong influencer of the bioavailability and binding of important ligands. Hemostasis Is Regulated by Force. Hemostasis is dependent on ECM. Exposure of underlying ECM to blood after disruption of a blood vessel’s endothelial lining begins the coagulation cascade, resulting in hemostasis and provisional matrix formation.103 Force is also involved in hemostasis and formation of provisional matrix after an injury. Von Willebrand factor (vWF) is important to hemostasis and is secreted as a multimer of many vWF proteins. These multimers remain bound to the endothelial cells from which they are secreted, or circulate in the blood and bind to collagen when ECM is exposed after an injury. Attachment to endothelial cells or binding to collagen allows the blood flow in a vessel to exert a shear force on the protein. This causes the protein to become more extended, and if the force is high enough (>21 pN), vWF’s binding site for platelet’s GPIbα is activated. The activation most likely occurs due to breakage of a hydrogenbonded fold, which exposes the GPIbα binding site in vWF’s A1 domain.104 Binding between vWF and GPIbα is an important early part of hemostasis, leading to platelet activation, formation of a platelet plug, and formation of the provisional matrix, since platelets become trapped in a polymerized network of fibrin. The length of vWF multimers regulates hemostasis, and the length is regulated by force. Longer vWF multimers experience more tensile force (proportional to the number of monomers squared) and, therefore, more activation of GPIbα binding sites. Force applied to multimers can unfold vWF’s A2 domain, exposing a site that is cleaved by ADAMTS13. The cleavage of multimers regulates their length and therefore the amount of flow required to activate binding to platelets.105 Mechanical control of hemostasis makes sense, since blood flow and mechanically supportive ECM structures are altered during an injury.

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CONCLUSIONS AND OUTLOOK The ECM is made up of many proteins interacting in complex ways. Cells use force to assemble matrix, and interactions between binding proteins control the structure and organization of the matrix, which ultimately determines the mechanical properties of the matrix. Many methods are available to generate ECM fibers in vitro; however, there has been limited characterization of how well these methods recapitulate the properties of cell-assembled ECM. Additionally, some ECM proteins have yet to be incorporated into in vitro-generated ECM fibers. Methods have been developed to measure the mechanical properties of ECM structures ranging from single nanofibers in the piconewton force range to bulk ECM structures in the kilonewton range. ECM composition and conformation can reveal information about the health of the tissue. Modeling can be used to make predictions about the structural mechanisms that allow stretch and unfolding to occur. ECM assembly and remodeling are regulated by force, as is ligand binding that impacts cell behavior or interaction with the ECM. Several of these mechanosensitive interactions have been discovered; it seems likely that there are many more cryptic binding sites and force-regulated interactions that have not yet been discovered. A major limitation to discovering mechano-biochemical interactions is a lack of highthroughput methods for identifying them. Additionally, there is a lack of methods to study these interactions in vivo to observe mechano-biochemical regulation in physiological H


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Biochemistry

(12) Carlisle, C. R., Coulais, C., Namboothiry, M., Carroll, D. L., Hantgan, R. R., and Guthold, M. (2009) The mechanical properties of individual, electrospun fibrinogen fibers. Biomaterials 30, 1205−1213. (13) Guthold, M., Liu, W., Sparks, E. A., Jawerth, L. M., Peng, L., Falvo, M., Superfine, R., Hantgan, R. R., and Lord, S. T. (2007) A Comparison of the Mechanical and Structural Properties of Fibrin Fibers with Other Protein Fibers. Cell Biochem. Biophys. 49, 165−181. (14) Zhong, C., Chrzanowska-Wodnicka, M., Brown, J., Shaub, A., Belkin, A. M., and Burridge, K. (1998) Rho-mediated Contractility Exposes a Cryptic Site in Fibronectin and Induces Fibronectin Matrix Assembly. J. Cell Biol. 141, 539−551. (15) Mouw, J. K., Ou, G., and Weaver, V. M. (2014) Extracellular matrix assembly: a multiscale deconstruction. Nat. Rev. Mol. Cell Biol. 15, 771−785. (16) Wenstrup, R. J., Florer, J. B., Brunskill, E. W., Bell, S. M., Chervoneva, I., and Birk, D. E. (2004) Type V Collagen Controls the Initiation of Collagen Fibril Assembly. J. Biol. Chem. 279, 53331− 53337. (17) Chen, S., and Birk, D. E. (2013) The regulatory roles of small leucine-rich proteoglycans in extracellular matrix assembly. FEBS J. 280, 2120−2137. (18) Ma, X., Schickel, M. E., Stevenson, M. D., Sarang-Sieminski, A. L., Gooch, K. J., Ghadiali, S. N., and Hart, R. T. (2013) Fibers in the Extracellular Matrix Enable Long-Range Stress Transmission between Cells. Biophys. J. 104, 1410−1418. (19) Abhilash, A. S., Baker, B. M., Trappmann, B., Chen, C. S., and Shenoy, V. B. (2014) Remodeling of Fibrous Extracellular Matrices by Contractile Cells: Predictions from Discrete Fiber Network Simulations. Biophys. J. 107, 1829−1840. (20) Tamariz, E., Grinnell, F., and Beckerle, M. (2002) Modulation of Fibroblast Morphology and Adhesion during Collagen Matrix Remodeling. Mol. Biol. Cell 13, 3915−3929. (21) Ingber, D. E. (2006) Cellular mechanotransduction: putting all the pieces together again. FASEB J. 20, 811−827. (22) Chen, C. S., and Ingber, D. E. (1999) Tensegrity and mechanoregulation: from skeleton to cytoskeleton. Osteoarthritis and Cartilage 7, 81−94. (23) Canty, E. G., Lu, Y., Meadows, R. S., Shaw, M. K., Holmes, D. F., and Kadler, K. E. (2004) Coalignment of plasma membrane channels and protrusions (fibripositors) specifies the parallelism of tendon. J. Cell Biol. 165, 553−563. (24) Canty, E. G., Starborg, T., Lu, Y., Humphries, S. M., Holmes, D. F., Meadows, R. S., Huffman, A., O’Toole, E. T., and Kadler, K. E. (2006) Actin Filaments Are Required for Fibripositor-mediated Collagen Fibril Alignment in Tendon. J. Biol. Chem. 281, 38592− 38598. (25) Kapacee, Z., Richardson, S., Lu, Y., Starborg, T., Holmes, D., Baar, K., and Kadler, K. (2008) Tension is required for fibripositor formation. Matrix Biol. 27, 371−375. (26) Broder, C., Arnold, P., Vadon-Le Goff, S., Konerding, M. A., Bahr, K., Muller, S., Overall, C. M., Bond, J. S., Koudelka, T., Tholey, A., Hulmes, D. J. S., Moali, C., and Becker-Pauly, C. (2013) Metalloproteases meprin and meprin are C- and N-procollagen proteinases important for collagen assembly and tensile strength. Proc. Natl. Acad. Sci. U. S. A. 110, 14219−14224. (27) Chakravarti, S., Magnuson, T., Lass, J. H., Jepsen, K. J., LaMantia, C., and Carroll, H. (1998) Lumican Regulates Collagen Fibril Assembly: Skin Fragility and Corneal Opacity in the Absence of Lumican. J. Cell Biol. 141, 1277−1286. (28) Giraud-Guille, M. M., Mosser, G., and Belamie, E. (2008) Liquid crystallinity in collagen systems in vitro and in vivo. Curr. Opin. Colloid Interface Sci. 13, 303−313. (29) Hohenester, E., and Yurchenco, P. D. (2013) Laminins in basement membrane assembly. Cell Adhesion & Migration 7, 56−63. (30) Sazonova, O. V., Isenberg, B. C., Herrmann, J., Lee, K. L., Purwada, A., Valentine, A. D., Buczek-Thomas, J. A., Wong, J. Y., and Nugent, M. A. (2015) Extracellular matrix presentation modulates vascular smooth muscle cell mechanotransduction. Matrix Biol. 41, 36−43.

and pathophysiological contexts. If force-regulated interactions are discovered to play an important role in disease, the binding partners involved could be used as targets for diagnostics or therapies. Force-regulated interactions discovered as part of normal development or tissue maintenance could also potentially be used to enhance tissue engineering and provide another knob to tune cell behavior and interactions.

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AUTHOR INFORMATION

Corresponding Author

*Phone: (617) 358-5489. Fax: (617) 353-6766. E-mail: [email protected]. ORCID

Gwendolyn A. Hoffmann: 0000-0001-7492-4503 Joyce Y. Wong: 0000-0002-3526-6381 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS G.A.H. was funded by the NIH Training Program in Quantitative Biology and Physiology (NIGMS 5T32 GM008764), the BUnano Cross-Disciplinary Fellowship, and the NIH Training Program in Cardiovascular Biology (NHLBI T32HL007969-15).

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ABBREVIATIONS ECM, extracellular matrix; Fn, fibronectin; BM, basement membrane; GAG, glycosaminoglycan; SLRP, small leucine rich proteoglycan; VEGF, vascular endothelial growth factor; TGFβ1, transforming growth factor beta 1; LOX, lysyl oxidase

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REFERENCES

(1) Frantz, C., Stewart, K. M., and Weaver, V. M. (2010) The extracellular matrix at a glance. J. Cell Sci. 123, 4195−4200. (2) Smith, E. M., Mitsi, M., Nugent, M. A., and Symes, K. (2009) PDGF-A interactions with fibronectin reveal a critical role for heparan sulfate in directed cell migration during Xenopus gastrulation. Proc. Natl. Acad. Sci. U. S. A. 106, 21683−21688. (3) Midwood, K. S., Williams, L. V., and Schwarzbauer, J. E. (2004) Tissue repair and the dynamics of the extracellular matrix. Int. J. Biochem. Cell Biol. 36, 1031−1037. (4) Kubow, K. E., Vukmirovic, R., Zhe, L., Klotzsch, E., Smith, M. L., Gourdon, D., Luna, S., and Vogel, V. (2015) Mechanical forces regulate the interactions of fibronectin and collagen I in extracellular matrix. Nat. Commun. 6. DOI: 10.1038/ncomms9026 (5) Hung, B. P., Hutton, D. L., and Grayson, W. L. (2013) Mechanical control of tissue-engineered bone. Stem Cell Res. Ther. 4, 10. (6) Wipff, P.-J., Rifkin, D. B., Meister, J.-J., and Hinz, B. (2007) Myofibroblast contraction activates latent TGF-β1 from the extracellular matrix. J. Cell Biol. 179, 1311−1323. (7) Ortiz Franyuti, D., Mitsi, M., and Vogel, V. (2018) Mechanical Stretching of Fibronectin Fibers Upregulates Binding of Interleukin-7. Nano Lett. 18, 15−25. (8) Zollinger, A. J., and Smith, M. L. (2017) Fibronectin, the extracellular glue. Matrix Biol. 60−61, 27−37. (9) Martino, M. M., and Hubbell, J. A. (2010) The 12th−14th type III repeats of fibronectin function as a highly promiscuous growth factor-binding domain. FASEB J. 24, 4711−4721. (10) Lo, C. M., Wang, H. B., Dembo, M., and Wang, Y. L. (2000) Cell movement is guided by the rigidity of the substrate. Biophys. J. 79, 144−152. (11) Engler, A. J., Sen, S., Sweeney, H. L., and Discher, D. E. (2006) Matrix Elasticity Directs Stem Cell Lineage Specification. Cell 126, 677−689. I


Perspective

Biochemistry (31) Bignon, M., Pichol-Thievend, C., Hardouin, J., Malbouyres, M., Brechot, N., Nasciutti, L., Barret, A., Teillon, J., Guillon, E., Etienne, E., Caron, M., Joubert-Caron, R., Monnot, C., Ruggiero, F., Muller, L., and Germain, S. (2011) Lysyl oxidase-like protein-2 regulates sprouting angiogenesis and type IV collagen assembly in the endothelial basement membrane. Blood 118, 3979−3989. (32) György, B., Tóth, E., Tarcsa, E., Falus, A., and Buzás, E. I. (2006) Citrullination: A posttranslational modification in health and disease. Int. J. Biochem. Cell Biol. 38, 1662−1677. (33) Vassiliadis, E., Oliveira, C. P., Alvares-da-Silva, M. R., Zhang, C., Carrilho, F. J., Stefano, J. T., Rabelo, F., Pereira, L., Kappel, C. R., Henriksen, K., Veidal, S. S., Vainer, B., Duffin, K. L., Christiansen, C., Leeming, D. J., and Karsdal, M. (2012) Circulating levels of citrullinated and MMP-degraded vimentin (VICM) in liver fibrosis related pathology. Am. J. Transl Res. 4, 403−414. (34) Kagan, H. M., and Li, W. (2003) Lysyl oxidase: Properties, specificity, and biological roles inside and outside of the cell. J. Cell. Biochem. 88, 660−672. (35) Cox, T. R., Bird, D., Baker, A.-M., Barker, H. E., Ho, M. W.-Y., Lang, G., and Erler, J. T. (2013) LOX-Mediated Collagen Crosslinking Is Responsible for Fibrosis-Enhanced Metastasis. Cancer Res. 73, 1721−1732. (36) Eyre, D. R., and Wu, J.-J. (2005) in Collagen (Brinckmann, J., Notbohm, H., and Müller, P. K., Eds.), pp 207−229. Springer: Berlin, Germany. (37) van der Slot, A. J., Zuurmond, A.-M., van den Bogaerdt, A. J., Ulrich, M. M. W., Middelkoop, E., Boers, W., Karel Ronday, H., DeGroot, J., Huizinga, T. W. J., and Bank, R. A. (2004) Increased formation of pyridinoline cross-links due to higher telopeptide lysyl hydroxylase levels is a general fibrotic phenomenon. Matrix Biol. 23, 251−257. (38) Griffin, M., Casadio, R., and Bergamini, C. M. (2002) Transglutaminases: nature’s biological glues. Biochem. J. 368, 377−396. (39) Chen, J. S. K., and Mehta, K. (1999) Tissue transglutaminase: an enzyme with a split personality. Int. J. Biochem. Cell Biol. 31, 817−836. (40) Wang, Z., Collighan, R. J., Gross, S. R., Danen, E. H. J., Orend, G., Telci, D., and Griffin, M. (2010) RGD-independent Cell Adhesion via a Tissue Transglutaminase-Fibronectin Matrix Promotes Fibronectin Fibril Deposition and Requires Syndecan-4/2 and α5β1 Integrin Co-signaling. J. Biol. Chem. 285, 40212−40229. (41) Chiquet, M., Birk, D. E., Bönnemann, C. G., and Koch, M. (2014) Collagen XII: Protecting bone and muscle integrity by organizing collagen fibrils. Int. J. Biochem. Cell Biol. 53, 51−54. (42) Marturano, J. E., Arena, J. D., Schiller, Z. A., Georgakoudi, I., and Kuo, C. K. (2013) Characterization of mechanical and biochemical properties of developing embryonic tendon. Proc. Natl. Acad. Sci. U. S. A. 110, 6370−6375. (43) Stern, R., Asari, A., and Sugahara, K. (2006) Hyaluronan fragments: An information-rich system. Eur. J. Cell Biol. 85, 699−715. (44) Almond, A. (2007) Hyaluronan. Cell. Mol. Life Sci. 64, 1591− 1596. (45) Kiani, C., Chen, L., Wu, Y. J., Yee, A. J., and Yang, B. B. (2002) Structure and function of aggrecan. Cell Res. 12, 19−32. (46) Wight, T. N. (2002) Versican: a versatile extracellular matrix proteoglycan in cell biology. Curr. Opin. Cell Biol. 14, 617−623. (47) Kinahan, M. E., Filippidi, E., Köster, S., Hu, X., Evans, H. M., Pfohl, T., Kaplan, D. L., and Wong, J. (2011) Tunable Silk: Using Microfluidics to Fabricate Silk Fibers with Controllable Properties. Biomacromolecules 12, 1504−1511. (48) Carlisle, C. R., Coulais, C., and Guthold, M. (2010) The mechanical stress−strain properties of single electrospun collagen type I nanofibers. Acta Biomater. 6, 2997−3003. (49) Jacobsen, M. M., Li, D., Gyune Rim, N., Backman, D., Smith, M. L., and Wong, J. Y. (2017) Silk-fibronectin protein alloy fibres support cell adhesion and viability as a high strength, matrix fibre analogue. Sci. Rep. 7, 45653. (50) Cao, L., Nicosia, J., Larouche, J., Zhang, Y., Bachman, H., Brown, A. C., Holmgren, L., and Barker, T. H. (2017) Detection of an

Integrin-Binding Mechanoswitch within Fibronectin during Tissue Formation and Fibrosis. ACS Nano 11, 7110−7117. (51) Ott, H. C., Matthiesen, T. S., Goh, S.-K., Black, L. D., Kren, S. M., Netoff, T. I., and Taylor, D. A. (2008) Perfusion-decellularized matrix: using nature’s platform to engineer a bioartificial heart. Nat. Med. 14, 213−221. (52) Saeidi, N., Karmelek, K. P., Paten, J. A., Zareian, R., DiMasi, E., and Ruberti, J. W. (2012) Molecular crowding of collagen: A pathway to produce highly-organized collagenous structures. Biomaterials 33, 7366−7374. (53) Little, W. C., Smith, M. L., Ebneter, U., and Vogel, V. (2008) Assay to mechanically tune and optically probe fibrillar fibronectin conformations from fully relaxed to breakage. Matrix Biol. 27, 451− 461. (54) Lu, H., Hoshiba, T., Kawazoe, N., and Chen, G. (2011) Autologous extracellular matrix scaffolds for tissue engineering. Biomaterials 32, 2489−2499. (55) Haynl, C., Hofmann, E., Pawar, K., Förster, S., and Scheibel, T. (2016) Microfluidics-Produced Collagen Fibers Show Extraordinary Mechanical Properties. Nano Lett. 16, 5917−5922. (56) Bradshaw, M. J., and Smith, M. L. (2019) Fibronectin fiber creep under constant force loading. Acta Biomater. 88, 78. (57) Backman, D. E., LeSavage, B. L., and Wong, J. Y. (2017) Versatile and inexpensive Hall-Effect force sensor for mechanical characterization of soft biological materials. J. Biomech. 51, 118−122. (58) Rohatgi, A. (2018) WebPlotDigitizer. https://automeris.io/ WebPlotDigitizer. (59) Sauer, F., Oswald, L., Ariza de Schellenberger, A., Tzschätzsch, H., Schrank, F., Fischer, T., Braun, J., Mierke, C. T., Valiullin, R., Sack, I., and Käs, J. A. (2019) Collagen networks determine viscoelastic properties of connective tissues yet do not hinder diffusion of the aqueous solvent. Soft Matter 15, 3055−3064. (60) Elbjeirami, W. M., Yonter, E. O., Starcher, B. C., and West, J. L. (2003) Enhancing mechanical properties of tissue-engineered constructs via lysyl oxidase crosslinking activity. J. Biomed. Mater. Res. 66A, 513−521. (61) Callister, W. D. (2007) Materials science and engineering: an introduction, 7th ed., John Wiley & Sons, New York. (62) Kesava Reddy, G., and Enwemeka, C. S. (1996) A simplified method for the analysis of hydroxyproline in biological tissues. Clin. Biochem. 29, 225−229. (63) Coulson-Thomas, V., and Gesteira, T. (2014) Dimethylme thylene Blue As say (DMMB). B IO-PROTOCOL 4. DOI: 10.21769/BioProtoc.1236 (64) Whitley, C. B., Ridnour, M. D., Draper, K. A., Dutton, C. M., and NeglIa, J. P. (1989) Diagnostic Test for Mucopolysaccharidosis. I. Direct Method for Quantifying Excessive Urinary Glycosaminoglycan Excretion. Clin. Chem. 35. (65) Naba, A., Clauser, K. R., Hoersch, S., Carr, S. A., Hynes, R. O., et al. (2012) The Matrisome: In Silico Definition and In Vivo Characterization by Proteomics of Normal and Tumor Extracellular Matrices. Mol. Cell. Proteomics 11, 18. (66) Angel, P. M., Baldwin, H. S., Gottlieb Sen, D., Su, Y. R., Mayer, J. E., Bichell, D., and Drake, R. R. (2017) Advances in MALDI imaging mass spectrometry of proteins in cardiac tissue, including the heart valve. Biochim. Biophys. Acta, Proteins Proteomics 1865, 927−935. (67) Cazares, L. H., Troyer, D. A., Wang, B., Drake, R. R., and John Semmes, O. (2011) MALDI tissue imaging: from biomarker discovery to clinical applications. Anal. Bioanal. Chem. 401, 17−27. (68) Kong, J., and Yu, S. (2007) Fourier Transform Infrared Spectroscopic Analysis of Protein Secondary Structures. Acta Biochim. Biophys. Sin. 39, 549−559. (69) Greenfield, N. J. (1999) Applications of circular dichroism in protein and peptide analysis. TrAC, Trends Anal. Chem. 18, 236−244. (70) Plaxco, K. W., Spitzfaden, C., Campbell, I. D., and Dobson, C. M. (1996) Rapid refolding of a proline-rich all-,8-sheet fibronectin type III module. Proc. Natl. Acad. Sci. U. S. A. 93, 10703. (71) Jakobsen, R., Brown, L., Hutson, T., Fink, D., and Veis, A. (1983) Intermolecular interactions in collagen self-assembly as J


Perspective

Biochemistry revealed by Fourier transform infrared spectroscopy. Science 220, 1288−1290. (72) Blackburn, P. R., Xu, Z., Tumelty, K. E., Zhao, R. W., Monis, W. J., Harris, K. G., Gass, J. M., Cousin, M. A., Boczek, N. J., Mitkov, M. V., Cappel, M. A., Francomano, C. A., Parisi, J. E., Klee, E. W., Faqeih, E., Alkuraya, F. S., Layne, M. D., McDonnell, N. B., and Atwal, P. S. (2018) Bi-allelic Alterations in AEBP1 Lead to Defective Collagen Assembly and Connective Tissue Structure Resulting in a Variant of Ehlers-Danlos Syndrome. Am. J. Hum. Genet. 102, 696−705. (73) Ruberti, J. W., and Hallab, N. J. (2005) Strain-controlled enzymatic cleavage of collagen in loaded matrix. Biochem. Biophys. Res. Commun. 336, 483−489. (74) Chen, X., Nadiarynkh, O., Plotnikov, S., and Campagnola, P. J. (2012) Second harmonic generation microscopy for quantitative analysis of collagen fibrillar structure. Nat. Protoc. 7, 654−669. (75) Wang, H.-W., Le, T. T., and Cheng, J.-X. (2008) Label-free imaging of arterial cells and extracellular matrix using a multimodal CARS microscope. Opt. Commun. 281, 1813−1822. (76) Tillberg, P. W., Chen, F., Piatkevich, K. D., Zhao, Y., Yu, C.-C., English, B. P., Gao, L., Martorell, A., Suk, H.-J., Yoshida, F., DeGennaro, E. M., Roossien, D. H., Gong, G., Seneviratne, U., Tannenbaum, S. R., Desimone, R., Cai, D., and Boyden, E. S. (2016) Protein-retention expansion microscopy of cells and tissues labeled using standard fluorescent proteins and antibodies. Nat. Biotechnol. 34, 987−992. (77) Cao, L., Zeller, M. K., Fiore, V. F., Strane, P., Bermudez, H., and Barker, T. H. (2012) Phage-based molecular probes that discriminate force-induced structural states of fibronectin in vivo. Proc. Natl. Acad. Sci. U. S. A. 109, 7251−7256. (78) Krammer, A., Craig, D., Thomas, W. E., Schulten, K., and Vogel, V. (2002) A structural model for force regulated integrin binding to fibronectin’s RGD-synergy site. Matrix Biol. 21, 139−147. (79) Lu, H., and Schulten, K. (1999) Steered molecular dynamics simulations of force-induced protein domain unfolding. Proteins: Struct., Funct., Genet. 35, 453−463. (80) Cavalcante, F. S. A., Ito, S., Brewer, K., Sakai, H., Alencar, A. M., Almeida, M. P., Andrade, J. S., Majumdar, A., Ingenito, E. P., and Suki, B. (2005) Mechanical interactions between collagen and proteoglycans: implications for the stability of lung tissue. J. Appl. Physiol. 98, 672−679. (81) Deravi, L. F., Su, T., Paten, J. A., Ruberti, J. W., Bertoldi, K., and Parker, K. K. (2012) Differential Contributions of Conformation Extension and Domain Unfolding to Properties of Fibronectin Nanotextiles. Nano Lett. 12, 5587−5592. (82) Salib, I. G., Kolmakov, G. V., Bucior, B. J., Peleg, O., Kröger, M., Savin, T., Vogel, V., Matyjaszewski, K., and Balazs, A. C. (2011) Using Mesoscopic Models to Design Strong and Tough Biomimetic Polymer Networks. Langmuir 27, 13796−13805. (83) Bradshaw, M. J., Cheung, M. C., Ehrlich, D. J., and Smith, M. L. (2012) Using Molecular Mechanics to Predict Bulk Material Properties of Fibronectin Fibers. PLoS Comput. Biol. 8, No. e1002845. (84) Meyers, M. A., and Chawla, K. K. (2008) Mechanical Behavior of Materials, 2nd ed., Cambridge University Press. (85) Chaudhuri, O., Gu, L., Darnell, M., Klumpers, D., Bencherif, S. A., Weaver, J. C., Huebsch, N., and Mooney, D. J. (2015) Substrate stress relaxation regulates cell spreading. Nat. Commun. 6. DOI: 10.1038/ncomms7365 (86) Bradshaw, M. J., and Smith, M. L. (2011) Contribution of Unfolding and Intermolecular Architecture to Fibronectin Fiber Extensibility. Biophys. J. 101, 1740−1748. (87) Li, W., Lucioni, T., Li, R., Bonin, K., Cho, S. S., and Guthold, M. (2017) Stretching single fibrin fibers hampers their lysis. Acta Biomater. 60, 264−274. (88) Paluch, E. K., Nelson, C. M., Biais, N., Fabry, B., Moeller, J., Pruitt, B. L., Wollnik, C., Kudryasheva, G., Rehfeldt, F., and Federle, W. (2015) Mechanotransduction: use the force(s). BMC Biol. 13. DOI: 10.1186/s12915-015-0150-4 (89) Malek, A. M. (1999) Hemodynamic Shear Stress and Its Role in Atherosclerosis. JAMA 282, 2035.

(90) Abu-Lail, N. I., Ohashi, T., Clark, R. L., Erickson, H. P., and Zauscher, S. (2006) Understanding the elasticity of fibronectin fibrils: Unfolding strengths of FN-III and GFP domains measured by single molecule force spectroscopy. Matrix Biol. 25, 175−184. (91) Ingham, K. C., Brew, S. A., Huff, S., and Litvinovich, S. V. (1997) Cryptic Self-association Sites in Type III Modules of Fibronectin. J. Biol. Chem. 272, 1718−1724. (92) Lemmon, C. A., Ohashi, T., and Erickson, H. P. (2011) Probing the Folded State of Fibronectin Type III Domains in Stretched Fibrils by Measuring Buried Cysteine Accessibility. J. Biol. Chem. 286, 26375− 26382. (93) Little, W. C., Schwartlander, R., Smith, M. L., Gourdon, D., and Vogel, V. (2009) Stretched Extracellular Matrix Proteins Turn Fouling and Are Functionally Rescued by the Chaperones Albumin and Casein. Nano Lett. 9, 4158−4167. (94) Oberhauser, A. F., Marszalek, P. E., Erickson, H. P., and Fernandez, J. M. (1998) The molecular elasticity of the extracellular matrix protein tenascin. Nature 393, 181. (95) Nemes, C., Ramsden, J. J., and Rozlosnik, N. (2002) The unfolding of native laminin investigated by atomic force microscopy. Phys. A 313, 578−586. (96) Camp, R. J., Liles, M., Beale, J., Saeidi, N., Flynn, B. P., Moore, E., Murthy, S. K., and Ruberti, J. W. (2011) Molecular Mechanochemistry: Low Force Switch Slows Enzymatic Cleavage of Human Type I Collagen Monomer. J. Am. Chem. Soc. 133, 4073− 4078. (97) Zareian, R., Church, K. P., Saeidi, N., Flynn, B. P., Beale, J. W., and Ruberti, J. W. (2010) Probing Collagen/Enzyme Mechanochemistry in Native Tissue with Dynamic, Enzyme-Induced Creep. Langmuir 26, 9917−9926. (98) Bhole, A. P., Flynn, B. P., Liles, M., Saeidi, N., Dimarzio, C. A., and Ruberti, J. W. (2009) Mechanical strain enhances survivability of collagen micronetworks in the presence of collagenase: implications for load-bearing matrix growth and stability. Philos. Trans. R. Soc., A 367, 3339−3362. (99) Chabria, M., Hertig, S., Smith, M. L., and Vogel, V. (2010) Stretching fibronectin fibres disrupts binding of bacterial adhesins by physically destroying an epitope. Nat. Commun. 1, 135. (100) Hubbard, B., Buczek-Thomas, J. A., Nugent, M. A., and Smith, M. L. (2016) Fibronectin Fiber Extension Decreases Cell Spreading and Migration: FIBRONECTIN FIBER EXTENSION DECREASES CELL SPREADING AND MIGRATION. J. Cell. Physiol. 231, 1728− 1736. (101) Buscemi, L., Ramonet, D., Klingberg, F., Formey, A., SmithClerc, J., Meister, J.-J., and Hinz, B. (2011) The Single-Molecule Mechanics of the Latent TGF-β1 Complex. Curr. Biol. 21, 2046−2054. (102) Klingberg, F., Chow, M. L., Koehler, A., Boo, S., Buscemi, L., Quinn, T. M., Costell, M., Alman, B. A., Genot, E., and Hinz, B. (2014) Prestress in the extracellular matrix sensitizes latent TGF-β1 for activation. J. Cell Biol. 207, 283−297. (103) Guyton, A. C., and Hall, J. E. (2006) Textbook of medical physiology, 11th ed., Elsevier Saunders: Philadelphia, PA. (104) Fu, H., Jiang, Y., Yang, D., Scheiflinger, F., Wong, W. P., and Springer, T. A. (2017) Flow-induced elongation of von Willebrand factor precedes tension-dependent activation. Nat. Commun. 8. DOI: 10.1038/s41467-017-00230-2 (105) Zhang, X., Halvorsen, K., Zhang, C.-Z., Wong, W. P., and Springer, T. A. (2009) Mechanoenzymatic Cleavage of the Ultralarge Vascular Protein von Willebrand Factor. Science 324, 1330−1334.

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