During migration, cells interact with their environment by exerting mechanical forces on it. A combination of two techniques shows that they do so in all three dimensions by a push–pull mechanism.
Mechanobiology is an emerging field that investigates how living cells sense and respond to the mechanical cues of their surroundings. In contrast to passive objects such as water droplets, living cells actively probe their environment by exerting forces on it as they migrate1. Such forces not only drive mechanical events such as cell deformation but also trigger cellular processes such as cell–environment adhesion signalling and cytoskeletal reorganization. In this context, mechanical forces have been shown1,2,3 to have a key role in many biological functions, including cell migration, cancer progression and stem-cell differentiation. But the precise characterization of these forces in space and time has remained elusive. Writing in Physical Review Letters, Delanoë-Ayari and colleagues4 describe a microscopic technique that does just that.
In the early 1980s, seminal work by Harris et al.5 demonstrated that cells can exert forces on and deform compliant two-dimensional (2D) substrates. Since then, various techniques have been developed to map the deformation induced by traction forces exerted by cells on elastic substrates1. These traction-force microscopy techniques have led to a greater understanding6,7,8 of the processes that regulate cell–substrate interactions, from the molecular to the multicellular level. However, until recently, the techniques have been used only to compute in-plane (horizontal) forces, thus assuming that the cellular biomechanical components responsible for the establishment of forces within cells were mostly oriented parallel to the surface. In other words, they presumed that the component of the forces that is perpendicular to the substrate was negligible (Fig. 1a).
However, we have come to learn that cells act in all dimensions as they probe and respond to the three-dimensional (3D) geometry of their environment9,10. In their study, Delanoë-Ayari et al.4 devise a method that can accurately map the 3D force pattern generated by adherent cells. The method is essentially an extension of the traction-force microscopy technique developed by Dembo and Wang11. It consists of measuring a cell's traction forces, and so a substrate's deformation, on the basis of the substrate's elastic properties and the displacement that the cell induces on fluorescent beads embedded near the substrate's surface. Combined with a technique12 that permits 3D tracking of the beads' dynamics, the method enables the spatial and temporal distributions of the cell's traction forces to be precisely determined in all directions.
The authors apply their method to cells of the soil-living amoeba Dictyostelium discoideum on a soft-gel substrate with easily controlled mechanical properties. Surprisingly, although the fluorescent beads are randomly distributed inside the gel, when focusing light on the gel's upper surface, the researchers observe a 'black hole' in the fluorescent signal just where the cell is located. This happens because the cell pushes the beads towards the gel's interior, causing them to go out of focus. The fluorescent signal re-emerges when the cell is removed from the substrate, because the beads recover their equilibrium position. What's more, the observed 3D force pattern clearly indicates that D. discoideum cells regulate their interactions with the soft substrate through a push–pull force mechanism: the cells push the gel vertically in the region underneath the cell nucleus but pull it obliquely towards the cell centre at the cell's edges (Fig. 1b). Because the overall force has to be zero, the pulling forces in the vertical direction exactly balance the pushing forces.
Delanoë-Ayari et al.4 demonstrate not only that vertical forces exist, but also that they are of the same order of magnitude as horizontal forces, thus highlighting the need to consider vertical forces in studies that examine the role of cell–substrate interactions in biological functions. Taken together with previous studies13,14 on mammalian tissue cells that showed that they deform their environment in much the same way as D. discoideum does, the authors' findings not only highlight the importance of taking into account 3D forces for all adherent cell types, but also give a new and clear description of the mechanical balance between the pushing and pulling forces.
Because substrate elasticity can govern cell fate3, one of the main issues in the field of mechanobiology concerns understanding the interplay between gene expression and mechanical forces exerted by cells on the environment. The observed force pattern raises questions about the physical coupling between the nucleus and the elastic components of the cytoplasm15. The authors cultured cells on substrates that are softer than the cells' nuclei, so the implication is that, on contact with the substrate, the cells deformed the substrate more than their nuclei were deformed. The question of whether, on stiffer substrates, the pushing forces could lead to nuclear deformation and cell-fate reprogramming requires investigation.
It is well known that on stiffer substrates mammalian tissue cells exert larger forces and are more spread out across the substrate surface, thus leading to a higher nuclear compression (Fig. 1c). Increasing substrate rigidity may therefore result in an increase in the horizontal forces and a relative decrease in the vertical ones. To what extent the authors' technique can be applied over a broad range of substrate stiffness remains an open question.
Although we are still far from a complete understanding of the mutual interaction between cell function and mechanical cues, Delanoë-Ayari et al. have shown that cellular traction forces in all three dimensions matter, and should be taken into account to fully understand cell–substrate interactions.