With the list of proteins encoded by the genome growing ever larger, a major challenge is to understand how they contribute to intracellular motility and cell movements. Historically, knowledge of motility has been built on a largely biochemical and structural foundation. The major biochemical components of the actin cytoskeleton have been identified, and their roles in defining the structure of the cytoskeleton and regulation are being discovered. However, motility is a dynamic process in which function is also built on the foundation of mechanics.
Dynamics implies a change in structure with time and includes movements of proteins and protein structures. From Newton's laws of physics, movements are linked with forces, but we are only now becoming capable of studying forces at the molecular and cellular levels. Much has changed from the pioneering studies of muscle physiologists who measured muscle fiber contractile forces with tension gauges and of experimental cell physiologists who estimated the contractile force of the contractile ring from the bending of glass needles. These pioneering studies lead directly to modern studies that incorporate elegant physical techniques such as optical traps capable of measuring piconewton forces or that deduce stiffness from images of microtubules and microfilaments buckling against an unmovable object. At the heart of these discoveries lies the capability of the light microscope to image single molecules and to watch the binding of individual ligands and enzyme action on individual nucleotides.
But to understand fully how a cell moves, we must be able to measure forces in the cell. In vivo methods that measure force directly are not yet invented. What is needed? A mechanical sensor, perhaps engineered from a fluorescent protein, that can change fluorescence when a force is applied. Such a technique would be the analog of experiments in which fluorescently labeled actin was microinjected into a live cell and scientists watched for the first time the actin cy-toskeleton treadmill at the leading edge of the cell.
A second challenge is to understand how a cell moves in its natural environment—between layers of cells in a three-dimensional extracellular matrix. The story of cell motility is one shaped by studies of cells moving on a flat surface. The flat world imposes a geometry in which interactions with the substratum is through adhesions on the ventral surface of the cell while the free, noninteracting surface defines the dorsal surface. Consider, instead, movement through a gel of matrix. When surrounded by matrix, cells lose their stereotypic flat shape and take on tubular spindle shape. With adhesions on all sides, cells corkscrew through the gel of matrix molecules. The adhesions are smaller than those formed with a flat surface, which itself makes studying adhesions more difficult. Furthermore, the techniques for estimating the forces that a cell must exert to crawl on a flat surface cannot be applied to a three-dimensional situation. Squeezing through a gel adds several dimensions of complexity to our understanding how cell motility and adhesion are coupled.
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