Cell migration: a tug-of-war inside your body

Original article: Physical forces during collective cell migration

If you ever played tug-of-war in elementary school, you might remember that it isn’t the friendliest game. People fall over, hands get burned from holding on to the rope, and knees get scraped from falling on the ground. Although victory can be sweet, the injuries that come with it may make you never want to play the game again. Perhaps surprisingly, there is a similar ‘’tug-of-war” happening inside your body, as individual cells move around from one place to another in a process called cell migration. What’s more, this microscopic tug-of-war may help to heal those scrapes and bruises that happened in elementary school, and those that happen in your everyday life.

A single cell moves by detaching and reattaching from the substrate, or the surface it is on, as the cell  expands and contracts. This movement exerts forces on the substrate. (These forces can actually be measured directly –  this is the topic of a previous softbites post.) When many cells move together in a “cell sheet”,  their motion becomes more complicated. Not only do cells push and pull on the substrate, but they also push and pull on the cells that surround them.  In today’s study, Xavier Trepat and colleagues show that there is a “tug-of-war” between cells that causes them to migrate.

Previously, it was thought that only the cells at the very front of the mass of migrating cell, or the leading edge of the cell sheet, exert forces on the substrate. According to this picture, most of the cells get passively pulled along by the leading edge, and neither push nor pull on the substrate. By measuring the forces the cells exert on the substrate, Trepat and his colleagues discovered that, in fact, all of the cells are involved in pushing the cell sheet forward.

The researchers measured the forces in a moving sheet of cells, taken from canine kidneys, growing on a gel substrate using a technique called traction force microscopy. The first step of this technique is to track the displacements of different points within the substrate as the cells move. Then, the mechanical properties of the gel are used to calculate the forces on the substrate generated by this motion. The researchers mapped the value of these forces using different colors, with red and blue representing very strong forces and black representing zero force. They first looked at what happened at the leading edge of the cell sheet, as in Figure 1.


Figure 1. a. Image of the cell sheet, in which individual cells are outlined in white. The field of view is 700 microns by 700 microns. b. The forces that the cells exert perpendicular to the leading edge of the cell sheet. c. The forces that the cells exert parallel to the edge of the cell sheet. Bright red and blue colors indicate strong forces (up to 100 Pa of stress), while black color indicates low forces. (Images adapted from the original article.) The cell sheet’s expansion was recorded in a video as well.

The researchers separated the normal forces (Figure 1b) — those exerted by the cells perpendicular to the leading edge of the cell sheet, or in the direction of the cells’ motion — from the forces exerted parallel to the leading edge of the cell sheet (Figure 1c). The bright red and blue colors in Figure 1 show that cells well inside the cell sheet exert forces on the substrate. From this, they hypothesized that instead of having “follower” and “leader” cells, all the cells contribute into pushing and pulling the cell sheet as they move.

The researchers then looked at larger areas of the cell sheet, such as that  shown in Figure 2. The bright colors near the edges correspond to strong forces,  while the black spots show that the forces in the center of the cell sheet are weaker. This suggests that the cell sheet “tugs” both to the right and the left as it expands. As the cells exert forces on the substrate, they exert forces on each other. The cells pulling to the right and the left are similar to two teams pulling a rope in a game of tug of war. The sheet of cells is like a rope that grows in the direction of the tugging of the cells.

Figure 2. Forces exerted by a larger piece of the cell sheet. Bright red indicates strong positive forces and blue indicates strong negative forces, while black indicates low forces. The scale bar on the bottom right is 200 micrometers.  (Image adapted from the original article.)

Next, the researchers wanted to understand how being tugged on by its neighbors affects the motion of individual cells: does the tug of war consistently pull a cell in a particular direction? Or is the cell equally likely to be pulled in any direction?  To answer this question, Trepat and colleagues measured the average force exerted on a cell by its neighbors, as a function of the distance of that cell from the edge of the sheet. If each cell was moving independently, the average normal force inside the sheet would be zero – on average, no cell would be pushing or pulling any other cell to a specific direction. Instead, as shown in Figure 3, the average force was not zero, and was actually higher for distances farther from the sheet’s leading edge. In other words, the cell sheet is expanding from the inside more than it’s being pulled from the edge.


Figure 3. The average normal force exerted on a cell by its neighbors, \sigma_{xx}, is higher farther from the leading edge of the cell sheet. (Figure adapted from the original article.)

Each individual cell crawling on a substrate has little effect on its surroundings, but many cells acting together can exert forces on each other to guide the collective in a particular direction. As cells replicate, such as in a healing wound, this guiding helps the cells expand in directions where there is space to be filled. This study by Trepat and colleagues reveals for the first time the tug-of-war that allows the tissues in our bodies to grow and heal.

The Peter Parker cell

Berginski M, Vitriol E, Hahn K, Gomez S [CC BY 2.5 (https://creativecommons.org/licenses/by/2.5)], via Wikimedia Commons

Original paper: Measuring mechanical tension across vinculin reveals regulation of focal adhesion dynamics


“USE YOUR LEGS!” That’s what might have been yelled at you the first time you went climbing. We are so used to walking or running that we don’t even think about how we do it. But when we face a new environment, such as a steep slope, we realize that finding the best strategy to move through space is not so easy. Now, imagine you are as small as few dozens of microns, without legs or arms, and you live in a viscous fluid. How would you move? This is the question biologists who are interested in cell movements have been trying to solve. By observing cells under a microscope, they saw that depending on their type or their environment, cells exhibit a wide variety of motion strategies. However, one thing never changes: cells need to exert forces on their environment to move. To do so, some kinds of cells create structures called focal adhesions. These structures are made up of several proteins, assembled on the outside of the cell. Like tiny bits of double-sided tape, their purpose is to stick the cell to whatever is nearby (see Figure 1). In slightly more technical language, focal adhesions connect the molecular skeleton of the cell to a substrate.

Figure 1. Movie of a moving cell with fluorescently labelled focal adhesions (from Berginski et al. 2011)

Cells can exert forces on their environment through focal adhesions. While it is possible to measure these forces outside the cell by engineering some force-sensing substrate [1], it is much trickier to understand what happens inside the cell. Accessing these forces inside the cells is the challenge Grasshoff and colleagues tackled in their 2010 paper.

In order to measure a force, the most straightforward method is to use a spring. A spring is a stretchable object for which, after calibration, we can relate its extension to the applied force. Therefore, a force can be measured by measuring the length of the spring. To measure the forces focal adhesions apply on the cell, one would need to inject tiny springs in the cells and connect them to the exerting-force structures.

To do this, the authors had the idea of taking advantage of a silk protein, produced by a spider, which is literally a molecular spring. Thanks to genetic tools, a part of the gene of this silk protein could be inserted within a gene called vinculin. The vinculin gene produces a protein that is an essential part of the focal adhesion structure. As shown in Figure 2A, vinculin connects the protein filaments of the cell skeleton to the outside of the cell (the extracellular matrix). The researchers engineered an artificial variant of vinculin that includes a molecular spring, derived from the silk protein, right in the middle of the naturally occurring vinculin molecule (see Figure 2B).

Figure 2. A. Schematic of focal adhesion. B. Schematic of the modified vinculin under low and high tension. Under high tension, the molecular spring is stretched. Red: adhesion protein, orange: vinculin head domain, yellow: vinculin tail domain, grey: contractile filaments. Arrows represent the magnitude of the tension.


After verifying that cells that are genetically modified to include the engineered focal adhesion protein behave normally, the next step was to measure the molecular spring extension. However, measuring distances at the molecular scale is not a piece of cake. For instance, the typical extension of such a spring is 6 nanometers, which is, by far, below the resolution of the best optical microscopes [2]. To circumvent this limitation, Grasshoff and colleagues took advantage of the Förster resonance energy transfer (FRET) effect to measure the distance between the two vinculin domains. The FRET effect takes place between two fluorescent molecules very close in space. A fluorescent molecule, when excited by a light at a precise wavelength, emits a light at a longer wavelength. But if a second fluorescent molecule is close enough, the first molecule (the donor) can directly transfer its energy to the second molecule (the acceptor). Then, the acceptor will emit light at an even larger wavelength than the donor’s. Consequently, the FRET intensity can be computed by measuring the relative emissions of the donor and acceptor molecules: the closer the acceptor is to the donor, the more energy the acceptor will absorb and re-emit. Furthermore, and importantly for this application, the efficiency of this process is very sensitive to the distance between the donor and the acceptor As a result, the distance between the two molecules can be measured with great precision (sub-nanometer) by measuring the intensity of the FRET effect. Therefore, the authors further engineered the vinculin protein by placing the molecular spring between two fluorescent molecules (Figure 3, yellow and red circles) that were capable of undergoing the FRET effect to measure the extension of the molecular spring.

Figure 3. Förster resonance energy transfer (FRET) effect in the modified vinculin of a focal adhesion under low and high tension. The excitation light of the donor molecule (yellow circle) is shown in green and the emission light of the acceptor molecule (red circle) is shown in red.

At this point, the authors had a method for measuring the tension intensity across vinculin molecules just by looking at the FRET intensity. In this way, they could generate a tension map  across the contacts of the cell with its environment. They saw that focal adhesion under high tension leads to a growth of the size of the focal adhesion which relieves it from its high tension. Perhaps surprisingly, they also showed that regions where the contact is extending (protruding areas) are under higher tension than regions where the contact is receding (retracting areas), as shown in Figure 4.

In this paper, the authors developed a new technique to measure forces inside cells. By conducting single-molecule experiments, they even could calibrate their engineered molecular spring and relate the FRET intensity to absolute values of forces (in the order of a few piconewtons [3]), paving the way to a whole class of new FRET-based force sensors with different stiffnesses, which can now be used in other structures inside cells.

Everything started with adding a spider silk gene in a cell. Such mutant cells have the amazing power of shading light on the cellular force machinery. But “with great power, comes great responsibility” as another spider mutant has once been told.

Figure 4. The FRET index (ratio of donor to acceptor fluorescence) reveals the state of tension through vinculin across a cell. Close-ups retracting areas (R1 and R2) show a high FRET index, ie. a low tension, and protruding areas (P1 and P2) show a low FRET index, ie. a high tension (adapted from Grashoff et al.).

[1] These techniques are called traction force microscopy. The deformation of calibrated substrate (either a gel or micropillars) is measured to calculate the forces exerted by the cell.
[2] Classical optical microscopes have a typical resolution of around 200 nm. New techniques of super-resolution microscopy reach a resolution of a few dozens of nanometers.
[3] To give you a sense of this order of magnitude, when you hold a pen of, let’s say 10 g, you apply a force of 0.1 N. At the cellular level, cells exert on their environment forces in the order of dozens of nanonewtons (according to this study). At the molecular level, DNA has been manipulated applying forces in the same range as the vinculin tension: 1-100 pN (according to this study).