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Arterial stiffness and inflammation are associated with atherosclerosis, and each have individually been shown to increase endothelial monolayer tension and permeability. The objective of this study was to determine if substrate stiffness enhanced endothelial monolayer tension and permeability in response to inflammatory cytokines. Porcine aortic endothelial cells were cultured at confluence on polyacrylamide gels of varying stiffness and treated with either tumor necrosis factor-α (TNFα) or thrombin. Monolayer tension was measured through vinculin localization at the cell membrane, traction force microscopy, and phosphorylated myosin light chain quantity and actin fiber colocalization. Cell permeability was measured by cell-cell junction confocal microscopy and a dextran permeability assay. When treated with TNFα or thrombin, endothelial monolayers on stiffer substrates showed increased traction forces, vinculin at the cell membrane, and vinculin phosphorylation, suggesting elevated monolayer tension. Interestingly, VE-cadherin shifted toward a smaller molecular weight in endothelial monolayers on softer substrates, which may relate to increased VE-cadherin endocytosis and degradation. Phosphorylated myosin light chain colocalization with actin stress fibers increased in endothelial monolayers treated with TNFα or thrombin on stiffer substrates, indicating elevated cell monolayer contractility. Endothelial monolayers also developed focal adherens intercellular junctions and became more permeable when cultured on stiffer substrates in the presence of the inflammatory cytokines. Whereas each of these effects was likely mitigated by Rho/ROCK, Rho/ROCK pathway inhibition via Y27632 disrupted cell-cell junction morphology, showing that cell contractility is required to maintain adherens junction integrity. These data suggest that stiff substrates change intercellular junction protein localization and degradation, which may counteract the inflammation-induced increase in endothelial monolayer tension and thereby moderate inflammation-induced junction loss and associated endothelial monolayer permeability on stiffer substrates.

The interplay between cell-cell and cell-substrate interactions is complex yet necessary for the formation and healthy functioning of tissues. The same mechanosensing mechanisms used by the cell to sense its extracellular matrix also play a role in intercellular interactions. We used the discrete element method to develop a computational model of a deformable cell that includes subcellular components responsible for mechanosensing. We modeled a three-dimensional cell pair on a patterned (two-dimensional) substrate, a simple laboratory setup to study intercellular interactions. We explicitly modeled focal adhesions and adherens junctions. These mechanosensing adhesions matured, becoming stabilized by force. We also modeled contractile stress fibers that bind the discrete adhesions. The mechanosensing fibers strengthened upon stalling. Traction exerted on the substrate was used to generate traction maps (along the cell-substrate interface). These simulated maps are compared to experimental maps obtained via traction force microscopy. The model recreates the dependence on substrate stiffness of the tractions' spatial distribution, contractile moment of the cell pair, intercellular force, and number of focal adhesions. It also recreates the phenomenon of cell decoupling, in which cells exert forces separately when substrate stiffness increases. More importantly, the model provides viable molecular explanations for decoupling: mechanosensing mechanisms are responsible for competition between different fiber-adhesion configurations present in the cell pair. The point at which an increasing substrate stiffness becomes as high as that of the cell-cell interface is the tipping point at which configurations that favor cell-substrate adhesion dominate over those favoring cell-cell adhesion. This competition is responsible for decoupling. ispartof: Biophysical Journal vol:119 issue:2 ispartof: location:United States status: Published online

We have developed a method to understand the forces that pairs of cells exert upon each other on a surface. When exerting a force or torque on another object, a cell experiences an equal and opposite recoil force, and must therefore brace itself against its substrate. If the substrate is soft and elastic, then absorbing the recoil causes observable displacements of marker particles imbedded below its surface. We here describe a method of traction force microscopy, which can detect and analyze the recoil displacements. We validate our method using a clone of Breast Ductal Epithelial cells, (BDE's), that crawl and interact on collagen coated polyacrylamide substrates of different stiffness. For these cells, noise in the bead data causes random, or “phantom”, forces that fluctuate with standard deviation of approximately 10 nN. In cases where cells are touching, we universally observe significant tension forces that are usually aligned with the centroid-centroid axis of the cell pair. In contrast there are only a few isolated cases where shear forces or torques between BDE cells are above threshold. We also observe that the BDE's exert tension forces in two very different ways; first as part of a “grappling” process which brings the cell membranes into greater contact, and second as the basis for a “tug-of-war,” which has the opposite result. Grappling seems to happen only in the early stages as BDE's pull themselves into alignment. Later on, during the tug-of-war, the cell-cell tension grows and can reach peak values of over 200 nN. In addition, the cell-substrate adhesions near the contact zone are lost.

There is now considerable evidence of the importance of mechanical cues in neuronal development and regeneration. Motivated by the difference in the mechanical properties of the tissue environment between the peripheral (PNS) and central (CNS) nervous systems, we compare substrate-stiffness-dependent outgrowth and traction forces from PNS (dorsal root ganglion (DRG)) and CNS (hippocampal) neurons. We show that neurites from DRG neurons display maximal outgrowth on substrates with a Young's modulus of ∼1000 Pa, whereas hippocampal neurite outgrowth is independent of substrate stiffness. Using traction force microscopy, we also find a substantial difference in growth cone traction force generation, with DRG growth cones exerting severalfold larger forces compared with hippocampal growth cones. The traction forces generated by DRG and hippocampal growth cones both increase with increasing stiffness, and DRG growth cones growing on substrates with a Young's modulus of 1000 Pa strengthen considerably after 18–30 h. Finally, we find that retrograde actin flow is almost three times faster in hippocampal growth cones than in DRG. Moreover, the density of paxillin puncta is significantly lower in hippocampal growth cones, suggesting that stronger substrate coupling of the DRG cytoskeleton is responsible for the remarkable difference in traction force generation. These findings reveal a differential adaptation of cytoskeletal dynamics to substrate stiffness in growth cones of different neuronal types, and highlight the potential importance of the mechanical properties of the cellular environment for neuronal navigation during embryonic development and nerve regeneration.

Extracellular matrices determine cellular fate decisions through the regulation of intracellular force and stress. Previous studies suggest that matrix stiffness and ligand anchorage cause distinct signaling effects. We show herein how defined noncovalent anchorage of adhesion ligands to elastic substrates allows for dissection of intracellular adhesion signaling pathways related to matrix stiffness and receptor forces. Quantitative analysis of the mechanical balance in cell adhesion using traction force microscopy revealed distinct scalings of the strain energy imparted by the cells on the substrates dependent either on matrix stiffness or on receptor force. Those scalings suggested the applicability of a linear elastic theoretical framework for the description of cell adhesion in a certain parameter range, which is cell-type-dependent. Besides the deconvolution of biophysical adhesion signaling, site-specific phosphorylation of focal adhesion kinase, dependent either on matrix stiffness or on receptor force, also demonstrated the dissection of biochemical signaling events in our approach. Moreover, the net contractile moment of the adherent cells and their strain energy exerted on the elastic substrate was found to be a robust measure of cell adhesion with a unifying power-law scaling exponent of 1.5 independent of matrix stiffness.

Cell adhesion and migration crucially depend on the transmission of actomyosin-generated forces through sites of focal adhesion to the extracellular matrix. Here we report experimental and computational advances in improving the resolution and reliability of traction force microscopy. First, we introduce the use of two differently colored nanobeads as fiducial markers in polyacrylamide gels and explain how the displacement field can be computationally extracted from the fluorescence data. Second, we present different improvements regarding standard methods for force reconstruction from the displacement field, which are the boundary element method, Fourier-transform traction cytometry, and traction reconstruction with point forces. Using extensive data simulation, we show that the spatial resolution of the boundary element method can be improved considerably by splitting the elastic field into near, intermediate, and far field. Fourier-transform traction cytometry requires considerably less computer time, but can achieve a comparable resolution only when combined with Wiener filtering or appropriate regularization schemes. Both methods tend to underestimate forces, especially at small adhesion sites. Traction reconstruction with point forces does not suffer from this limitation, but is only applicable with stationary and well-developed adhesion sites. Third, we combine these advances and for the first time reconstruct fibroblast traction with a spatial resolution of approximately 1 microm.

Conventional studies of dynamic phagocytic behavior have been limited in terms of spatial and temporal resolution due to the inherent three-dimensionality and small features of phagocytosis. To overcome these issues, we use a series of frustrated phagocytosis assays to quantitatively characterize phagocytic spreading dynamics. Our investigation reveals that frustrated phagocytic spreading occurs in phases and is punctuated by a distinct period of contraction. The spreading duration and peak contact areas are independent of the surface opsonin density, although the opsonin density does affect the likelihood that a cell will spread. This reinforces the idea that phagocytosis dynamics are primarily dictated by cytoskeletal activity. Structured illumination microscopy reveals that F-actin is reorganized during the course of frustrated phagocytosis. F-actin in early stages is consistent with that observed in lamellipodial protrusions. During the contraction phase, it is bundled into fibers that surround the cell and is reminiscent of a contractile belt. Using traction force microscopy, we show that cells exert significant strain on the underlying substrate during the contraction phase but little strain during the spreading phase, demonstrating that phagocytes actively constrict during late-stage phagocytosis. We also find that late-stage contraction initiates after the cell surface area increases by 225%, which is consistent with the point at which cortical tension begins to rise. Moreover, reducing tension by exposing cells to hypertonic buffer shifts the onset of contraction to occur in larger contact areas. Together, these findings provide further evidence that tension plays a significant role in signaling late-stage phagocytic activity.

Embryonic development depends on effective cell migration whose malfunction leads to abnormalities. Migration is the integrated outcome of a cycle of inter-connected component processes, namely protrusion, adhesion and contraction. Most, if not all, molecular details of these processes have been established. The major remaining challenge is to identify mechanisms that couple these processes in space and time. Compared to a relatively well-established interaction between protrusion and adhesion, however, there is no understanding as to how contraction and adhesion interact dynamically at the time scale of a single migration cycle, and whether these interactions affect protrusion through adhesion-protrusion coupling. The major hurdle that makes it challenging to investigate contraction- protrusion link is from technology: there is nearly no tool to quantify myosin II- based contraction in cytoskeletal network compared to numerous imaging approaches for characterization of protrusion - adhesion coupling. Here, we hypothesize that contraction dynamically modulates adhesion at a distance, which in turn promotes or inhibits protrusion via several redundant mechanical and signaling pathways. To test this hypothesis, we developed a continuum mechanical (CM) model to infer location and time of intracellular forces in migrating cells, which will be compared against high-resolution traction force microscopy (TFM) to obtain absolute force levels and infer material heterogeneity in the cytoskeleton. Preliminary results from Ptk1 cell wound-healing assay show that intracellular force field and traction force field are highly correlated, suggesting the feasibility of the absolute intracellular force level reconstruction. To establish the ‘information flow’ between contraction, adhesion and protrusion, we will use a correlation analysis of spontaneous fluctuations to show the coupling and information flow between them in unperturbed cells and in a cell where candidate molecules mediating the putative link between contraction and protrusion is slightly perturbed.

Cells sense and respond to their environment according to many factors, including gravity. Changes in the gravitational field during space exploration may alter cellular interactions. Our understanding of the fundamental mechanisms by which gravitational forces ultimately affect cell function, however, is limited. Based on our prior observations, human mesenchymal stem cells (hMSCs) adopt a more rounded morphology during simulated microgravity (clinorotation). We hypothesize that microgravity affects the cell-substrate forces, which in turn affects cell adhesion and motility. Therefore, we investigated the correlation between traction forces, spreading and chemotaxis. As an extension to our previously reported “clinochip” device, we developed a lab-on-a-chip device suitable for implementing traction force microscopy during clinorotation. The device contains a channel coated with an array of fluorescent beads embedded in a polyacrylamide substrate that can be processed to calculate cell-substrate traction forces. For our studies, we investigated both hMSCs and osteosarcoma cancer cells (143-B), because they represent highly regulated and deregulated cell states in the osteogenic lineage, respectively. Clinorotation speeds of 0, 30, and 75 rpm were examined, and cell shape, adhesion area, traction forces, and chemotactic migration were measured. Interestingly, results indicate that hMSCs exhibit a dose-dependent response to clinorotation speed based on a shift in the population distribution of cell shape and adhesion area, while osteosarcoma cells do not. These results suggest that a deregulated cell phenotype may possess distinct mechanosensing characteristics, which may be related to our measures of cell adhesion, traction and chemotaxis. Our results are among the first efforts to directly measure the physical interplay between the cell and its substrate during simulated microgravity. This will allow us to gain a deeper understanding of the cellular mechanisms that lead to tissue-level changes, such as atrophy and reduced bone mineral density, observed in astronauts.

Mechanical stimuli profoundly alter cell fate, yet the mechanisms underlying mechanotransduction remain obscure due to a lack of methods for molecular force imaging. In this presentation, I will describe the development of second and third generation mechanophores (molecules that fluoresce upon experiencing force) for imaging molecular forces exerted by individual integrin cell surface receptors. Specifically, we developed a new class of molecular tension probes that function as a switch to generate a 20-30-fold increase in fluorescence upon experiencing a threshold piconewton force. The probes employ immobilized DNA-hairpins with tunable force response thresholds, ligands, and fluorescence reporters. Quantitative imaging reveals that integrin tension is highly dynamic and increases with an increasing integrin density during adhesion formation. Mixtures of fluorophore-encoded probes show integrin mechanical preference for cyclized-RGD over linear-RGD peptides. Multiplexed probes with variable guanine-cytosine content within their hairpins reveal integrin preference for the more stable probes at the leading tip of growing adhesions near the cell edge. DNA-based tension probes are among the most sensitive optical force reporters to date, overcoming the force and spatial-resolution limitations of traction force microscopy. The application of these sensors to image forces associated with a range of mechano-regulatory processes that occur at the lipid membrane of the cell, such as endocytosis, Notch receptor activation, and integrin adhesion receptors will be described as well.

Forces exerted by stationary cells have been investigated on the level of single focal adhesions by combining elastic substrates, fluorescence labeling of focal adhesions, and the assumption of localized force when solving the inverse problem of linear elasticity theory. Data simulation confirms that the inverse problem is ill-posed in the presence of noise and shows that in general a regularization scheme is needed to arrive at a reliable force estimate. Spatial and force resolution are restricted by the smoothing action of the elastic kernel, depend on the details of the force and displacement patterns, and are estimated by data simulation. Corrections arising from the spatial distribution of force and from finite substrate size are treated in the framework of a force multipolar expansion. Our method is computationally cheap and could be used to study mechanical activity of cells in real time.

Hepatocellular carcinoma (HCC) is the fifth most common cause of cancer-related mortality over the world, and liver cirrhosis was reported as the most important risk factor for HCC development. However, the roles of mechanotransduction in HCC have not been fully explored yet. Kappa-actin (κ-actin), a novel class of actin correlated with poor postoperative survival, was selected as our study model. We investigated how κ-actins regulate HCC cells using stiffness adjustable polymeric matrixes, which mimic the cirrhotic microenvironment. Different cellular physiology, such as proliferation, contact topography and 3-dimensional invasion, were observed in microenvironment with different stiffness. Interestingly, with high k-actin expression, we noticed the significantly decrease of focal adhesions (FAs) formation while substrate's stiffness > 16 kPa. Photoactive microscopy showed lower stability of k-actin structures, and the unstable actin organization can contribute to the decrease of FAs/Adherence junction (AJs) formations. Furthermore, the instability of AJs (E-cadherin) may correlate to the observed higher invasion of k-HCC in vitro. Traction force microscopy (TFM) was developed to further quantify the mechanical interactions at the ECM-cell/cell-cell interfaces. By measuring the deformation of polymeric gel substrates, the force balance at the contact surfaces can be reconstructed. We observed the ECM-cell traction force increased with the increasing substrate stiffness, and the traction force decreased with the k-actin expression, which consisted with the spatial pattern of FAs. In summary, we showed the expression of k-actin alters the formation of FAs/AJs in HCC, which can contribute to the observed high cell proliferation and invasion of HCC. Furthermore, using photoactive microscopy, we demonstrated the actin dynamics play the essential roles in FAs and AJs formation of HCC. These findings may contribute to our understanding about the influence of actins on HCC through mechanotransduction perspective.

Mesenchymal stem cells (MSCs) have been known to exhibit substrate stiffness-dependent differentiation, and history of the mechanical dose from culture environment to MSCs sensitively is found to alter its phenotype. A certain level of substrate stiffness and duration period on that determine the fate of MSCs. In relation to this, we have found before that microelastically-patterned hydrogel with heterogeneous distribution of matrix stiffness allow MSCs to suppress fate determination into specific differentiation lineages, and contribute to keep the undifferentiated state. We call such mode of MSCs as “frustrated differentiation”, which serves to construct culture substrate for MSCs to maintain their stemness in high-qualified state. The basis of this phenomenon is in the enforced oscillation of mechanical dose from environment to MSCs during the nomadic migration between stiff and soft region of gel matrix, which eliminate the history of experience on a certain level of stiffness. To design such heterogeneous microelastically-patterned gels, we have applied the photolithographic microelasticity patterning using photoculable gelatins. The emergence of frustrated mode of differentiation was previously confirmed by immunofluorescence and RT-PCR analysis for the expression markers, but more detailed and precise characterizations are of course required. In this study, to fully characterize the frustrated differentiation of MSCs, we investigated oscillation of the mechanical dose and mechanical signal input to MSCs employing the long-term traction force microscopy for MSCs culture on the microelastically-striped patterned gels. In addition, we performed cDNA microarray analysis for the MSCs culture in such mode of frustrated differentiation. As the result, MSCs in normadic movement between stiff and soft region of gel surface were confirmed to exhibit characteristic transcriptome and marked large fluctuation profile of traction forces compared with plain control gels.

Cells organize actin filaments into higher-order structures by regulating the composition, distribution and concentration of actin crosslinkers. Palladin is an actin-crosslinking protein that is found in the lamellar actin network and stress fibers, two actin structures critical for mechanosensing of the physical environment. Palladin also serves as a molecular scaffold for alpha-actinin, a key actin crosslinker. By virtue of its close interactions with actomyosin structures in the cell, palladin may play an important role in cell mechanics. However, the role of palladin in cellular force generation and mechanosensing has not been studied. In this study we use human pancreatic tumor associated fibroblasts (TAFs) to investigate the role of palladin in regulating the plasticity of the actin cytoskeleton and cellular force generation in response to alterations in substrate stiffness. Traction force microscopy revealed that TAFs are sensitive to substrate stiffness as they generate larger forces on substrates of increased stiffness. Contrary to expectations, knocking down palladin increased the forces generated by cells, and also inhibited the ability to sense substrate stiffness for very stiff gels. This was accompanied by significant differences in the actin organization and adhesion dynamics of palladin knock down cells. Perturbation experiments also suggest altered myosin activity in palladin KD cells. Our results suggest that the actin crosslinkers such as palladin and myosin motors coordinate for optimal cell function and to prevent aberrant behavior as in cancer metastasis.

Generation of mechanical force is essential for the function of tissue cells, for example during migration, wound healing or rigidity sensing. The traction field of adherent cells can be measured on soft elastic substrates using traction force microscopy (TFM). However, mathematical reconstruction of traction fields effectively requires inversion of the long-ranged elastic equations and therefore is an ill-posed inverse problem. Moreover measurement of the traction pattern alone does not tell us how force is distributed inside the cell. To improve the robustness, resolution and scope of conventional TFM, we have developed a new procedure called model-based TFM (MBTFM). We estimate the distribution of intracellular tension from elastic substrate data by minimizing the difference between the experimentally measured displacement field and the predictions of a detailed theoretical model based on active cable networks. Previously this type of model has been successfully used to predict cell shapes on micropatterned substrates. For MBTFM, we consider not only active network contraction, but also contributions of various types of contractile bundles modeled as contractile one-dimensional line element embedded into the network. Subsequent computer simulations of network contraction and parameter optimization allow us to estimate the most likely distribution of tension over various contractile structures and adhesion sites.

Mechanical interactions between cell and substrate are involved in vital cellular functions from migration to signal transduction. A newly developed technique, traction force microscopy, makes it possible to visualize the dynamic characteristics of mechanical forces exerted by fibroblasts, including the magnitude, direction, and shear. In the present study such analysis is applied to migrating normal and transformed 3T3 cells. For normal cells, the lamellipodium provides almost all the forces for forward locomotion. A zone of high shear separates the lamellipodium from the cell body, suggesting that they are mechanically distinct entities. Timing and distribution of tractions at the leading edge bear no apparent relationship to local protrusive activities. However, changes in the pattern of traction forces often precede changes in the direction of migration. These observations suggest a frontal towing mechanism for cell migration, where dynamic traction forces at the leading edge actively pull the cell body forward. For H-ras transformed cells, pockets of weak, transient traction scatter among small pseudopods and appear to act against one another. The shear pattern suggests multiple disorganized mechanical domains. The weak, poorly coordinated traction forces, coupled with weak cell-substrate adhesions, are likely responsible for the abnormal motile behavior of H-ras transformed cells.

The mechanical respons of the micro environment of a cell is of great influence for its differentiation and mobility. However, the mechanism that a cell uses to sense these mechanical properties remains largely unkown. Mechanosensing could happen by direct coupling to the nucleus via the actin cytoskeleton or by activating molecular signaling cascades through protein complexes.A recent combination of traction force microscopy with super resolution imaging allowed for novel research into the nanoscale architecture of force-bearing focal adhesions. Here we apply this technique to investigate the relation between local force exertion by the cell and the number of signal transduction proteins inside the integrin adhesome.Integrin adhesomes are mechanosensory protein complexes that couple the intracelular force bearing actomyosin structures to the extracelular environment. These complexes contain signal transduction proteins like paxillin, vinculin, talin and FAK, that change conformation when force is exerted on the complex.By immunostaining these proteins with Alexa647 their position can be localized to 20nm in fixed cells. However, due to the photophysical properties of Alexa647, and the binding stoichiometry in antibody staining the number of localizations does not scale linearly with the number of signal transduction proteins. To overcome this problem we present a novel analysis method to relate the number of localizations to the minimal number of signal transduction proteins.

Directional cell migration is involved in a broad range of biological phenomena, ranging from the metastatic spreading of cancer to wound healing. Chemotaxing Dictyostelium cells adapt their morphology and speed to external conditions like the stiffness and adhesive properties of their substrate. The mechanism by which they control both their shape and speed remains largely unknown. Using Traction Force Microscopy measurements, we construct traction tension kymographs to examine the spatio-temporal dynamics of both the adhesions and the traction stresses during migration. We show that wild-type cells control their motility by switching between two motility modes with distinct adhesion and contractility dynamics. In the “Stepping-Stepping” mode, the adhesion sites remain stationary while the cell moves forward by periodic axial contractions. The back adhesions break after new frontal adhesions are formed. In the “Stepping-Gliding” mode, the cell reduces the magnitude of the traction stresses, increases the frequency of axial contractions and its migration speed, and keeps the frontal adhesion stationary while sliding the back adhesion forward. These two modes are not conserved when cells move on adhesive poly-L-Lys coated substrates, where cells alternate between a “Nearly Stationary” mode, characterized by strong lateral contractions and extremely low migration speed and a “Gliding-Gliding” mode, where multiple weak and transient adhesions are formed which are gliding forward as the cell moves by barely adhering to the substrate. In summary, our findings have contributed to a more precise understanding of how the coordination of traction stresses together with the adhesion dynamics result in efficient amoeboid cell migration. We propose that these are highly conserved mechanisms, which function in a range of amoeboid cells, including leukocytes, as well as other forms of cell motility.

The actin cytoskeleton is known to be a main structural and mechanical component of many eukaryotic cells. Dynamic crosslinking of actin filaments by proteins such as alpha-actinin, can change these mechanics, however, the effect of dynamic crosslinking on cell mechanics has not been previously reported. Here we quantify the viscoelastic moduli of cells with alpha-actinin isoforms that have varied binding affinity.We apply several techniques, including passive and active micro-rheology and AFM measurements to measure cellular moduli with alpha-actinin isoforms. The experiments are also performed after myosin inhibition and ATP depletion to study the contribution of active contractility to the viscoelastic properties of these cells. AFM-based stress relaxation and dynamic indentation tests are used to calculate the mechanical properties of the cellular cortex, while active micro-rheology using optical tweezers allows us to calculate the storage and loss moduli of the cytoplasm.Furthermore, the overall energy dissipation is estimated at cellular level, and these results are compared with cell work on the elastic substrates, using traction force microscopy data. Energy dissipation via dynamic crosslinkers in the cytoskeleton can regulate cellular viscoelasticity and basic biological functions such as spreading, force-generation, and migration.

The ability to differentiate human cardiomyocytes (heart muscle cells) from induced pluripotent stem cells (iPSCs) presents high potential to model heart contractility. Shortening of sarcomeres in series along intracellular myofibrils enables the beating of cardiomyocytes. We developed a platform with patterned single iPSC-cardiomyocytes in arrays and tested the contractility of these cells as a function of their shape and of substrate stiffness. We fluorescently labelled sarcomeres with LifeAct to analyze their shortening and organization. We measured the mechanical output of contractile cycles with traction force microscopy and adapted cross-correlation algorithms to characterize movement of sarcomeres. We assayed single cells on patterns of 2000 μm2 with aspect ratios (length:width) ranging from 1:1 to 7:1 and on substrates with varied stiffnesses: 6 kPa, 10 kPa and 35 kPa. Preliminary studies suggested that cell morphology and substrate stiffness affect the organization of sarcomeres with a potential impact in cardiomyocyte contractility. We aimed to understand how the contractility of iPSC-cardiomyocytes is affected by the improved organization of sarcomeres induced by these cues. For a substrate stiffness of 10 kPa, we observed that cells with a 7:1 aspect ratio produced higher contractile forces per μm of sarcomere shortening. Improved sarcomere alignment seemed to drive increased contractility of cells with this shape. Substrate stiffness also affected the contractility of 7:1 iPSC-cardiomyocytes. 35 kPa substrates induced sarcomere ruptures and lower contractile forces. Cells on 6 kPa substrates developed sarcomeres that buckled. These results suggested a role of intracellular tension in contractility. We further tested how tension affects contractility and sarcomere organization in 7:1 cells through cell stretching, culture in high calcium and by inhibiting non-muscle myosin. Results showed evidence that a tuned intracellular tension mechanism drives myofibril alignment and improved contractility in iPSC-cardiomyocytes.

Depending on cell type and the local environment, tumor cells show a variety of different migration modes, including mesenchymal and amoeboid motion. As the cell interacts with a spatially changing extracellular matrix, cell movements are highly heterogeneous in space and time and thus do not comply with conventional statistical models. By explicitly incorporating this heterogeneity into mathematical models, we are able to quantify how cells change their migration behavior over time. Based on Bayesian hierarchical modeling, the temporal evolution of directional persistence and migratory activity of each cell can be reconstructed from its measured migration path. We demonstrate that the temporal changes in persistence and activity provide a distinct fingerprint of the strategies that cells employ to cope with different environments. For example, persistence is positively correlated with activity in a 3D collagen matrix over much longer time periods compared to migration on 2D substrates, supporting the hypothesis that cells are able to pull themselves along collagen fibers and hence use the surrounding matrix to their advantage. To test this hypothesis, we measure cell pulling forces in a collagen gel with 3D traction force microscopy. We find that directional persistence of invading MDA-MB-231 breast carcinoma cells is highly correlated with contractility and cell elongation. The invasion behavior of these cells can thus be described by a gliding motion with alternating phases of simultaneously high or low contractility, elongation, migratory activity and persistence.

Adult mesenchymal stem cells (MSCs) “feel” the stiffness of their environment and differentiate in response to it; thus aberrant stiffness resulting from fibrosis in muscle dystrophies could misdirect MSCs into the wrong lineage. Conversely, MSCs have been shown to respond in a myosin-dependent manner to adipogenic and osteogenic media when cell spread area changes from 103 to 104 μm2 or when cultured in specific shapes, e.g. circles versus rectangles, and polygons, respectively. When both cues are present in a disparate fashion, e.g. highly elongated cells similar to muscle despite the presence of an abnormally stiff microenvironment, we hypothesized that a myosin contraction-dependent balance could induce a subset of MSCs to differentiate in to a muscle-like phenotype despite residing in a dystrophic-like stiffness. To regulate MSC morphology, we patterned fibronectin in shapes of varying aspect ratios but common area on polyacrylamide substrates of known stiffness. MSCs spread to the patterns and localized their focal adhesion in a stiffness and shape-dependent manner. Using traction force microscopy, we found that strain energy from cell-generated forces scaled with stiffness, but decreased as a function of cell elongation with isotropic cell patterns producing the highest contractile energy in contrast to our hypothesis. Muscle-specific myosin heavy chain (mMHC), an indicator of early muscle differentiation, also was expressed in a stiffness and elongation dependent manner. On muscle-like stiffness of 11 kiloPascals (kPa), cells with only minimal elongation, i.e. 1:1 and 3:1 patterns, expressed mMHC most strongly. In contrast on osteogenic-like matrices of 34 kPa, highest MHC expression corresponded to the most elongated patterns. These shape- and stiffness-dependent lineage changes with muscle markers correlated to contractility-based observations suggest that muscle induction may be possible in non-permissive stiffer environments and could prove beneficial to treat fibrotic muscle diseases.

Force generation is the main physiological function of primary heart muscle cells (myocytes). To study this experimentally, we cultivated primary myocytes from neonatal rat pups on silicone elastomer substrates of controlled elasticities. Young's moduli ranged from 1 to 500 kPa covering physiological and pathological heart tissue stiffnesses. Fluorescent microbeads embedded in the surface of these substrates served as markers for the displacements caused by spontaneously contracting cells. Forces of isolated cells were extracted from these displacements by the method of traction force microscopy (TFM). Transfection with GFP-α-actinin enabled visualization of cell adhesions simultaneously to TFM. By restricting the force application points chosen for the TFM algorithm to certain adhesion structures their mutual relevance for cellular traction force application was explored. Moreover, total cell forces clearly depended on substrate stiffness. These experiments were complemented by a quantification of myofiber contraction. Here, GFP-α-actinin containing z-bands of separate myofibers were imaged by fluorescence microscopy. Subsequent digital image processing was used to quantify the local contraction of individual fibers on different stiffnesses. In contrast to forces, fiber contraction appeared to depend little on substrate stiffness.

The effect that growth factors such as epidermal growth factor (EGF) have on cell-cell adhesion is of interest in the study of cellular processes such as epithelial-mesenchymal transition. Because cell-cell adhesions cannot be measured directly, we use three-dimensional traction force microscopy to measure the tractions applied by clusters of MCF-10A cells to a compliant substrate beneath them before and after stimulating the cells with EGF. To better interpret the results, a finite element model, which simulates a cluster of individual cells adhered to one another and to the substrate with linear springs, is developed to better understand the mechanical interaction between the cells in the experiments. The experiments and simulations show that the cluster of cells acts collectively as a single unit, indicating that cell-cell adhesion remains strong before and after stimulation with EGF. In addition, the experiments and model emphasize the importance of three-dimensional measurements and analysis in these experiments.

Tissue cells typically utilize their actomyosin contractile machinery to actively pull on their environment, which can be either the extracellular matrix or neighboring cells. The mechanical forces that a cell exerts and experiences have been shown to regulate fundamental cellular processes, including cell growth, proliferation, differentiation and migration. However, little is known about the spatial distribution of mechanical stress in tissues. In particular, the extent to which mechanical forces are communicated through cell-cell interactions across a tissue is not well understood.Here, we present a novel method, based on high resolution traction force microscopy, to measure mechanical stresses that are transmitted through cell-cell interfaces in small cellular clusters (∼10 cells). Cells are classified according to the number of neighboring cells. We find that this degree of cellular connectivity can determine many properties, including the amount of force transmitted through a particular cellular interface. In order to determine how force balance in the cell cluster is locally achieved, we compared forces transmitted through cells to forces exerted on the underlying substrate. A correlation analysis of these forces reveals the length scale over which forces can be transmitted through the cell cluster. Furthermore, by molecular perturbations, we are identifying proteins that may be essential for long range stress communication in the cluster. The ability to quantify force communication between cells will allow us to examine how cell-cell mechanical interactions contribute to overall tissue stress and vice versa. It will also allow us to investigate the role of mechanical stresses in establishing signaling gradients. This will further our understanding of the role of mechanical stress in processes that require fine coordination between cells, such as collective migration in morphogenesis and cancer.

Chemotaxis is involved in a broad range of biological phenomena such as during cancer metastasis. It requires a tightly regulated, spatiotemporal coordination of underlying biochemical processes that impact the mechanics of cell migration. In response to intrinsic and environmental cues, motile cells can adapt their migration effectively. Yet both the mechanisms by which this adaptation occurs and the role of the interactions between biochemistry and mechanics of cell migration are largely unknown.We use Fourier Traction Force Microscopy to measure the spatiotemporal evolution of shape and traction stresses and construct traction tension kymographs to analyze cell motility as a function of the dynamics of the cells' mechanically active adhesions (traction adhesions). We show that wild-type cells migrate mainly by forming two stationary traction adhesion sites at their front and back halves, over which the cell body moves forward in a step-wise fashion through periodic axial and, to a lesser degree, lateral contractions. We demonstrate that lateral forces are critical in mediating cell motility and essential for migration on highly adhesive substrates where cells implement two alternate motility modes to achieve migration. Our analysis of two mutant strains that lack distinct F-actin crosslinkers (mhcA- and abp120- cells) also supports a key role for lateral contractions in amoeboid cell motility, while the differences in their traction adhesion dynamics suggest the two mutant strains use distinct mechanisms to achieve migration. The considerable differences we find in both the spatiotemporal organization of traction adhesions and contractility, when comparing to the control wild type, provide insight into the role of the extracellular environment and of key cytoskeletal proteins in cell migration. We propose that these are highly conserved mechanisms, which function in a range of amoeboid cells, including leukocytes, as well as other forms of cell motility.

Traction force microscopy (TFM) allows imaging of the traction field exerted by a cell during adhesion and spreading on an elastic hydrogel. We used a combination of TFM and microfluidics to measure the traction forces and motility of human neutrophils under both chemokinesis and chemotaxis in response to formyl-met-leu-phe (fmlp). Using polyacyrlamide gels functionalized with intercellular adhesion molecule-1 (ICAM-1), we show that neutrophil traction stresses can be measured across a broad range of gel stiffnesses, from 6 to 20 kPa. We found neutrophil directed motion is caused by a rearward squeezing uropodial stress, and the cell motion is always counter to this motion; this is true both in chemokinesis as well as chemotaxis. During turning, the orientation of the rearward stress precedes turning. Cells exert larger forces in chemotaxis (r.m.s. force of ∼ 100pN) than in chemokinesis (∼ 50 pN). On surfaces of different compliance, cells move with a greater force and a higher chemotatic index on stiffer substrates; these changes occur without a change in neutrophil speed. In the same magnitude gradient, cells move directedly and with greater force if the mean concentration of chemoattractant is closer to the KD of receptor binding. Blocking with an antibody against β2-integrin (TS1/18) completely eliminates traction forces and directed motion. RhoA has been implicated as signal transduction agent in the cell that is responsible for rearward contractile stress and the direction of neutrophil motion; we show here that inhibition of a GTPase down stream of RhoA (ROCK) with a pharmacological agent reduces directional motion and force generation, and leads to abnormal morphology in which rearward contraction is compromised. Taken together, neutrophil directed motion and force generation result from an interplay between substrate adhesiveness and uropodial contractility through RhoA.

T cell activation is critical for the adaptive immune response in the body. The binding of the T cell receptor (TCR) with antigen on the surface of antigen presenting cells (APC) triggers signaling cascades and cell spreading. Physical forces exerted through the TCR have been shown to induce signaling events, but the origin of how these forces are generated and maintained is unknown. Here, we use traction force microscopy to measure the forces exerted by Jurkat T cells during TCR activation. We used anti-CD3 coated elastic polyacrylamide gels to stimulate Jurkat T cells and measured the spatially resolved traction stress map exerted by these cells as they were activated. Perturbation experiments revealed that stresses were largely generated by actin assembly and disassembly and regulated by the flow speed of actin. Our experiments further suggest that TCRs are structurally linked to the actin cytoskeleton through the Arp2/3 complex. On the other hand, we found that myosin II motor activity was dispensable for maintenance of traction stresses, but was important for traction stress generation. Finally, we investigated calcium influx in Jurkat T cells when activated on substrates of physiologically relevant stiffnesses. Our results highlight the importance of cytoskeletal forces for receptor activation in T cells.

Traction Force Microscopy (TFM) is a powerful approach for quantifying cell-material interactions that over the last two decades has contributed significantly to our understanding of cellular mechanosensing and mechanotransduction. In addition, recent advances in three-dimensional (3D) imaging and traction force analysis (3D TFM) have highlighted the significance of the third dimension in influencing various cellular processes. Yet irrespective of dimensionality, almost all TFM approaches have relied on a linear elastic theory framework to calculate cell surface tractions. Here we present a new high resolution 3D TFM algorithm which utilizes a large deformation formulation to quantify cellular displacement fields with unprecedented resolution. The results feature some of the first experimental evidence that cells are indeed capable of exerting large material deformations, which require the formulation of a new theoretical TFM framework to accurately calculate the traction forces. Based on our previous 3D TFM technique, we reformulate our approach to accurately account for large material deformation and quantitatively contrast and compare both linear and large deformation frameworks as a function of the applied cell deformation. Particular attention is paid in estimating the accuracy penalty associated with utilizing a traditional linear elastic approach in the presence of large deformation gradients.

Cell migration is a vital process in which cells undergo directed movement to particular locations. However, the molecular mechanisms that coordinate this process are still poorly understood. Here, two techniques are used in tandem to study cell migration. Spatio-Temporal Image Correlation Spectroscopy (STICS) can create vector maps of protein velocities inside a living cell while Traction Force Microscopy (TFM) allows us to measure the forces applied by a cell on a substrate, which play a crucial role in cell migration. Using these techniques together allows us to simultaneously compare adhesion protein velocities inside the cells and the underlying traction forces exerted by the cell, to provide greater insight into cell migration. We show that traction forces and retrograde paxillin flow exhibit directional correlation but an inverse magnitude relationship in motile U2OS cells, indicating slippage at the molecular clutch level. By coating polyacrylamide gels with different ligands, we observe differences in the strength of those integin-ligand binding pairs. Finally, a new correlation analysis is introduced that can differentiate between potential adhesion movement models.View Large Image | View Hi-Res Image | Download PowerPoint Slide

The traction force microscopy (TFM) is a technique widely used to measure cellular traction forces that are closely related to cell migration, mechanical signaling, and other cellular functions. We apply the TFM to characterize the dynamic force patterns in different crawling modes of Caenorhabditis elegans (C. elegans) on soft gel matrices of different stiffness. When C. elegans crawls forward, it concentrates the thrust force to localized regions along the body rather than forming a uniform load distribution in its lateral direction. The dynamic force distributions appeared differentially in different behavioral modes of C. elegans including the forward, backward movement, as well as a sharp turn called the Ω-turn. Such dynamic behaviors of C. elegans might be considered as an effort to minimize drag resistance by reducing contact area between its body and gel surface, and these observations are very similar to recent experimental study suggested for the slithering of snake on flat surface. This work was supported by the National Research Foundation (NRF) grant 2013-012420 (J. Park) and 2010-0016886 (S. Ahn and J. Shin).

Increasing evidence suggests mechanical forces modulate T cell function. In this report, we investigate forces applied by mouse CD4+ T cells onto an underlying substrate as a model of the interface between T cells and antigen presenting cells. Traction force microscopy was carried out using microfabricated arrays of elastomer (polydimethylsiloxane, Sylgard 184, PDMS) pillars; cells induce deflections of the pillars which can be measured and used to estimate force applied to each structure. We chose a pillar geometry of 1 micrometer diameter and 5-9 micrometer height. These pillars were coated with a 1:1 mix of activating antibodies to CD3 and CD28, which ligate and activate the TCR complex and costimulatory CD28 signal. Traction force microscopy carried out on mouse naive CD4+ T cells 1 hour after seeding revealed that naive cells exert forces onto these pillar arrays with magnitude on the order of 50 pN per structure. Moreover, force application by a given cell is periodic, with a cycle on the order of minutes. To investigate the physiological implications of these forces, we measured IL-2 secretion by T cells seeded onto planar PDMS substrates of varying rigidity, which was controlled by varying the ratio of base : curing agent, yielding bulk moduli of 2 MPa to 25 kPa. Substrates were coated with activating antibodies to CD3 and CD28 prior to cell seeding, and the per area concentration of antibodies varied less than 10% across the different moduli. Activation of naive T cells, measured as IL-2 secretion over six hours, was 50% greater on the stiffest vs. softest elastomers, and each condition was statistically different from all others (Kruskal-Wallis methods, alpha = 0.05). These results demonstrate a functional impact of mechanical forces on T cell activation, and reveal new dynamics of the immune synapse.

Cells change their form and function by assembling actin stress fibers at their base and exerting traction forces on their extracellular matrix (ECM) adhesions. Individual stress fibers are thought to be actively tensed by the action of actomyosin motors and to function as elastic cables that structurally reinforce the basal portion of the cytoskeleton; however, these principles have not been directly tested in living cells, and their significance for overall cell shape control is poorly understood. Here we combine a laser nanoscissor, traction force microscopy, and fluorescence photobleaching methods to confirm that stress fibers in living cells behave as viscoelastic cables that are tensed through the action of actomyosin motors, to quantify their retraction kinetics in situ, and to explore their contribution to overall mechanical stability of the cell and interconnected ECM. These studies reveal that viscoelastic recoil of individual stress fibers after laser severing is partially slowed by inhibition of Rho-associated kinase and virtually abolished by direct inhibition of myosin light chain kinase. Importantly, cells cultured on stiff ECM substrates can tolerate disruption of multiple stress fibers with negligible overall change in cell shape, whereas disruption of a single stress fiber in cells anchored to compliant ECM substrates compromises the entire cellular force balance, induces cytoskeletal rearrangements, and produces ECM retraction many microns away from the site of incision; this results in large-scale changes of cell shape (> 5% elongation). In addition to revealing fundamental insight into the mechanical properties and cell shape contributions of individual stress fibers and confirming that the ECM is effectively a physical extension of the cell and cytoskeleton, the technologies described here offer a novel approach to spatially map the cytoskeletal mechanics of living cells on the nanoscale.

The interplay between chemical and mechanical signals plays an important role in cell biology, and integrin receptors are the primary molecules involved in sensing and transducing external mechanical cues. We used integrin-specific probes in molecular tension fluorescence microscopy to investigate the pN forces exerted by integrin receptors in living cells. The molecular tension fluorescence microscopy probe consisted of a cyclic Arg-Gly-Asp-D-Phe-Lys(Cys) (cRGDfK(C)) peptide tethered to the terminus of a polyethylene glycol polymer that was attached to a surface through streptavidin-biotin linkage. A fluorescence resonance energy transfer mechanism was used to visualize tension-driven extension of the polymer. Surprisingly, we found that integrin receptors dissociate streptavidin-biotin tethered ligands in focal adhesions within 60 min of cell seeding. Although streptavidin-biotin binding affinity is described as the strongest noncovalent bond in nature, and is ∼106 - 108 times larger than that of integrin-RGD affinity, our results suggest that individual integrin-ligand complexes undergo a marked enhancement in stability when the receptor assembles in the cell membrane. Based on the observation of streptavidin-biotin unbinding, we also conclude that the magnitude of integrin-ligand tension in focal adhesions can reach values that are at least 10 fold larger than was previously estimated using traction force microscopy-based methods.

The mechanical properties of the cellular microenvironment regulate processes that include migration, proliferation, and differentiation. These interactions occur at adhesions, which serve as both traction points and signaling hubs and mediate bi-directional sensing and responses to specific features of the surrounding extracellular matrix (ECM). Adhesions execute these activities through an intricate network of putative molecular interactions that largely remain to be demonstrated and characterized functionally in living cells. The challenge is to capture the highly localized and transient associations that characterize these activities in adhesions and determine how they respond to different microenvironments. In this study, we use high-resolution fluorescence fluctuation microscopy to map the formation and stoichiometry of integrin-associated complexes in the adhesions that populate the leading edge of migrating cells. We focus on putative integrin activating (kindlin and talin) and actin-linking (talin, vinculin and a-actinin) molecules and show that all molecules are present in adhesions as soon as they are visible; however, they form integrin containing complexes hierarchically, at different times, with variable stoichiometry within the adhesion itself and change as the adhesion matures into larger structures. To parse out the effects of the mechanical properties of the ECM on the numbers, aggregation states, and associations of these molecules, we extend these measurements to substrates with variable stiffness and correlate them with high-resolution traction force microscopy (TFM) measurements. We show that individual and newly formed adhesions at the leading edge of protruding cells transmit forces on soft as well as stiff substrates, with force magnitudes that correlate with the integrated intensity of the adhesions and the total number of individual adhesion molecules. These measurements provide novel information on complex formation as adhesions evolve and respond to substrate rigidity.

Directed migration of endothelial cells is crucial for angiogenesis and vascular remodeling. This migration is known to depend on both chemical and mechanical interactions. It has been shown that HUVECs migrate towards VEGF, but individual cell tracking in a stable and quantifiable gradient has not been done. In order to control for both chemical and mechanical interactions, we have used a microfluidic device that can generate a chemical gradient of VEGF in a spatially and temporally stable manner. This microfluidic device is placed over a polyacrylamide gel so that a range of physiological substrates, on which HUVECs migrate, could be tested. HUVECs were individually tracked and observed to chemotax towards higher VEGF concentrations on a variety of substrates. We were also able to apply traction force microscopy to HUVEC migration inside these chemotactic microfluidic chambers and found that HUVECs generate more force when moving directionally rather than randomly. These results indicate that the chemical gradient as well as the substrate mechanics affect both HUVEC migration and force generation.

Focal adhesions (FAs) are micron-sized protein assemblies that coordinate cell adhesion, migration, and mechanotransduction. How the many proteins within FAs are organized into force sensing and transmitting structures is poorly understood. We combined fluorescent molecular tension sensors with super-resolution light microscopy to visualize traction forces within FAs with

The mechanical forces exerted and detected by living cells play integral roles in diverse biological phenomena, including growth and development, wound healing, and cancer metastasis. In the past decade, techniques such as traction force microscopy and micropost arrays have proven to be powerful tools for measuring the forces generated by cells. In particular, traction force microscopy has recently been extended to three-dimensional cell culture environments by embedding tracer beads in either a synthetic polyethylene glycol hydrogel (PEG; Legant et al., Nat. Meth. 2010) or in collagen gels (Koch et al., PLoS ONE 2012). The embedded beads move in response to cell-generated distortions of the matrix, allowing cell-generated forces to be calculated. We sought to develop an experimental system that would exhibit the excellent mechanical properties of the PEG hydrogel while using a naturally occurring biological matrix. Fibrin gels fulfill both of these requirements: fibrin is elastic up to ∼50% strain (Brown et al., Science 2009) and is also widely used for 3D cell culture. Here we describe the use of fluorescently labeled fibrin gels to measure the forces generated by cells in 3D culture. We observe dramatic but elastic deformations of the fibrin matrix surrounding cells as they grow, divide, and migrate. Further, we find that the dynamic forces generated by the cell can be measured using the deformations of the matrix itself, providing a direct observation of how the cell modifies its surroundings. We discuss the use of this new technique in studying matrix remodeling and cell migration.

Many cellular events require dynamic changes in the shape and structural organization of the cell morphology and presumably affect gene expression by changing the nuclear shape. In this work, we investigate the effect of cell shape on the nucleus and the mechanism by which intracellular forces are transmitted to the nucleus. We analyzed the intracellular reorganization of individual endothelial cells plated on micropatterned substrates, imposing cells to spread on various aspect ratios. Specific drugs were used to alter each component of the cytoskeleton and we observed the spatial reorganization of the actin network, microtubules, intermediate filaments and focal adhesions, as well as the nuclear shape. Our data demonstrate the key role of the actin cytoskeleton in the adaptation of nuclear shape with cell elongation. Indeed, we show that the nucleus is subjected to pincer forces generated within the cytoskeleton via actin stress fibers. These intracellular forces drastically affect the nuclear shape and decrease the nuclear volume by 40-50% before attaining a state that is highly resistant to further deformation. Based on the quantification of cell traction forces by traction force microscopy, we propose a mechanical model that accounts for our observation and quantitatively predicts the nuclear shape. Our work also demonstrates that nucleus adaptation to cell elongation leads to a modification of nuclear functions. Indeed, DNA staining reveals an increase in chromatin condensation in highly deformed and compressed nucleus. We show that nuclear deformation in response of cellular elongation results in a strong decrease of cell ability to enter S phase and thus to proliferate. In conclusion, our results demonstrate that the shape of the cell is transposed by the actin cytoskeleton to the nucleus and suggest that it can alter the accessibility of genes to the transcription machinery.

To understand cell adhesion and migration, it is crucial to have information on cellular traction stress. Most traction force microscopy only measures shear stress. Recently a few group developed develop methods to measure normal stress and mostly based on finite element method. We report our finite element-method to compute three-dimensional (3D) traction stress exerted by human mesenchymal stem cells on a two-dimensional compliant polyacrylamide substrate embedded with fluorescent nanoparticles. The images of nanoparticles are acquired by confocal microscopy. We tracked the displacement of nanoparticles and computed strain, stress and strain energy by ANSYS. We examined both experimental factors and calculation that affect the resolution of the force measurement, especially in z direction. The stress measurement of the third dimension provides complete information on traction stress.

The fusion of mammalian myoblast cells is an integral part of muscle growth and regeneration. Although the biochemical aspects of cell fusion have been extensively investigated, a quantitative description of the physical phenomena underlying this process has not yet been explored. This study aims to quantify and distinguish both the extracellular and intercellular forces generated by two myoblasts during fusion or the lack thereof. To achieve this goal, we fabricated a protein-patterned polyacrylamide hydrogel with embedded fluorescent particles to employ time-lapse traction force microscopy on isolated pairs of cells. Our results indicate the presence of large intracellular stresses due to the polarizing tangential traction vectors pointing away from the cell-cell interface in both fusing and non-fusing cells. For non-fusing cells, these polarizing stresses were maintained throughout the duration of the time-lapse microscopy as the cells shift to and fro on the protein patterns. However, for cells undergoing fusion, the polarization of the traction vectors ceases as the stresses generated by the cell injecting its nuclei dominates the other. In addition to the tangential forces, we also analyzed the particle displacement field for normal stresses, which are revealed to be negligible under all circumstances. The magnitudes of the normal stresses have been shown to exist at an order of magnitude lower than that of the tangential stresses.

The infiltration of circulating neutrophils into the vascular wall is an early step leading to inflammation and atherogenesis, and requires transendothelial migration and invasion through the basement membrane. While much research has been dedicated to the identification of the signaling cascade involved in neutrophil recruitment, less is known about how neutrophils generate the three-dimensional (3-D) forces and shape changes required for invasion. To address this issue, differentiated HL-60 neutrophil-like cells are plated on a reconstituted basement membrane made of Matrigel and incorporated with chemoattractant fMLP. The cells subsequently adhere and invade into the 3-D Matrigel. Using 3-D Fourier traction force microscopy, we measure the evolution of cell shape and traction stresses in this 3-D invasion model. Our results show that several filopodia-like cell extensions at cell edge lead the way and exert pulling forces, which contributes to generate a large invasive protrusion that burrows into the Matrigel by generating pushing forces. We find that the total pulling force from all the filopodia structures balances the pushing force exerted by the invasive structure, revealing a coordination between morphodynamic changes and 3-D traction stresses during cell invasion. Furthermore, since the number of filopodia exceeds the number of invasive protrusions, this force balance causes the pushing stresses created at the invasive site to be much higher than the pulling stresses exerted by the filopodia. Thus, the 3-D traction forces exerted by filipodia play an important role in regulating protrusion dynamics of invading neutrophils in 3-D microenvironments, and are mechanistically coordinated to enhance invasion.

Mechanical interactions between cells and the surrounding extracellular matrix, such as adhesion, contraction, and force transduction, play a central role in many fundamental cell behaviors, including proliferation, cell death, and motility. The ability to precisely manipulate the intracellular machinery that regulates these interactions could therefore provide a powerful tool for controlling the mechanical properties of living cells and could also allow us to re-engineer how cells sense and respond to mechanical stimuli in their microenvironment, which would be particularly useful for tissue engineering and cellular technologies where cells are interfaced with synthetic microenvironments. Towards this goal, we have genetically engineered stable cell lines in which we can precisely and dynamically alter the mechanobiological behavior of living cells by varying the activity of signal transduction proteins, such as RhoA GTPase, using constitutively active and dominant negative mutants under the control of a tetracycline-repressible promoter. Through a variety of imaging and biophysical techniques, including atomic force microscopy and traction force microscopy, we have demonstrated graded and dynamic control over cytoskeletal architecture, cell shape and spreading, contractility, and cellular stiffness. In addition, using glioblastoma multiforme as a model system, we show how these cell lines can be used to study the effects of altered cellular mechanical properties on cancer cell motility and invasion.

There are strong indications that mechanical forces are particularly relevant in immune recognition. For the immune system, the enormous variety of antigenic ligands imposes a fundamental challenge to the discriminative power; the mechanism for discriminating between activating and non-activating ligands has remained enigmatic.In a recent theoretical study we showed how forces alters the potency for receptor ligand discrimination by orders of magnitudes(1). For the T cell receptor, which specifically binds to peptides presented by MHC on an antigen-presenting cell, discrimination can be realized with kinetic proofreading, which fails when ligands have only marginal differences in their off-rates. We showed, however, that the specificity of antigen-recognition can be massively improved by putting the TCR-pMHC bond under load: while under no force the bond rupture probability decays exponentially with time, force-induced bond rupture leads to much narrower distributions.Here, we present cellular traction force microscopy data to measure forces involved during T cell activation. Hydrogels were prepared with variable stiffness. Fluorescent beads carrying CD3 antibodies were immobilized onto the top layer of the hydrogel. Forces are read out by measuring the fluorescent bead movement throughout T cell attachment and activation. The bead movement was directly correlated to forces applied to the antibodies immobilized on the beads. Moreover, discrimination between lateral and transversal applied forces was possible by tracking the beads' positions in 3D.1.Klotzsch, E., and G.J. Schutz. 2013. Improved Ligand Discrimination by Force-Induced Unbinding of the T Cell Receptor from Peptide-MHC. Biophysj. 104: 1670-1675.

Many important biological processes, such as endothelial mechanotransduction of hemodynamic forces and neutrophil extravasation, involve the transmission of stresses across a cell monolayer. During these processes, the monolayer undergoes both lateral distortion due to the in-plane traction forces generated by the cells, and bending due to the out-of-plane component of the traction forces. However, the contribution of this bending to the monolayer stresses has been neglected in the literature. Here, we present a novel technique to determine monolayer stresses that considers both lateral distortion and bending. To illustrate the method, and to quantify the relative importance of the lateral and bending stresses, we measure the monolayer stresses in micropatterned endothelial cell islands of varying sizes and shapes. The cell islands are cultured on flexible polyacrylamide gels embedded with fluorescent beads, which deform due to traction forces exerted by the cells. We measure the three-dimensional gel deformation using previously established 3D Traction Force Microscopy methods, and recover the monolayer stresses from the measured deformation using Kirchoff-Love thin plate theory. The equations are solved numerically in an efficient manner using a Fourier pseudo-spectral method. The boundary conditions corresponding to the geometry of the cell islands are enforced within the Fourier framework using a relaxation iterative method. Our results indicate that, regardless of island shape, the three-dimensional bending stresses are dominant at the center of the island while the lateral stresses are more important near the island edge. Also, comparing the results from islands of different sizes shows that the relative importance of the bending stresses decreases with island size. These results suggest that it is necessary to resolve bending stresses to accurately determine the monolayer stresses, and reveal that the transmission of forces across cell junctions is three-dimensional and more complex than previously believed.