Myosin II dynamics and the influence of S100A4

Cells contain many thousands of components that interact in a coordinated way to give rise to properties characteristic of life (e.g. cell division, motility). Myosin is a key protein component that polymerises to form filamentous structures that are part of the cell cytoskeleton that help both maintain and change cell shape. These filaments are dynamic and myosin components may associate and dissociate over a period of seconds, even under conditions where the overall filament appears to be stable for many minutes. The cell regulates the assembly of myosin filaments by a number of mechanisms including interactions with other proteins, of which S100A4 appears a significant factor. S100A4 is a small Ca2+ binding protein that attaches to the myosin tail and inhibits polymerisation. Changes in the concentration of S100A4 in cells can lead to changes in morphology and migration behaviour. The objectives of this proposal are to study the detailed time courses of binding between S100A4, myosin and other proteins in pure state and to devise microscope-based assays through which their interactions can be followed in living cells. Preliminary work by the applicants has established many of the tools required for this research. The atomic structure of S100A4 has been determined and its interaction sites on myosin have been partially characterised. A cell line has been developed whose S100A4 concentration can be changed at will through specific induction and the use of siRNA (a specific molecule which blocks the expression of this protein). Furthermore the myosin can be expressed as fusion with a green fluorescent protein so that the myosin filaments can be visualised under the microscope. A custom-built microscope has been developed to allow a thin section of the cell to be observed and within this area as small region can be photobleached by brief exposure to laser light. While this destroys the fluorescence of the fusion protein, the myosin part is unaffected and continues to associate and dissociate from filaments. The time course of recovery of the bleached area provides key information about the diffusion rates of the proteins and their exchange rates into filamentous structures. We will also change rates of interaction by modulating the intracellular Ca2+ concentration. This will be done using a ultraviolet light flash to breakdown an unstable complex of Ca2+ which is loaded into the cell by prior incubation. The interaction rates determined within the cell will be compared with those observed with purified proteins in solution in the absence of other cellular components to see if they agree. Absence of agreement will indicate other components that are involved and that will need to be defined. Because of the complexity of cellular interactions, careful controls will be required to determine if specific effects arise directly from myosin-S100A4 interactions. One approach is to make mutations in the myosin and S100A4 so their binding sites are destroyed. Knowledge of the atomic structure of S100A4 and the region within the myosin tail that it interacts with, will aid this approach. Further work is proposed to define the complete interaction region at high resolution. One difficulty in carrying out quantitative measurements on living cells is the variation in cell shape between samples. We will explore patterning techniques whereby a favourable substrate is deposited on a slide in various shapes (e.g. X, Y and U) which should encourage the cells to bind with a well-defined shape. Furthermore, the corners of the patterns will encourage the formation of adhesion complexes, while cytoskeletal structures such as a actomyosin stress fibres will be positioned between the extremities (e.g. X will induce a square-shaped cell with four sets of stress fibres linking each corner). Such immobilised cells should allow better reproducibility of measurements.

High-resolution structures of S100A4 and its binding partners (the C terminal rod of myosin IIA and liprin beta 1) will be characterised by crystallography and NMR methods. The interaction between these proteins will also be studied by transient-state kinetic methods. The influence of Ca2+ on the kinetics will be measured by perturbing ion concentrations using flash photolysis of caged Ca2+ and caged chelators. These data will be correlated with GFP-myosin IIA dynamics and the influence of S100A4 as measured by fluorescence recovery after photobleaching (FRAP) within A431 cells. Levels of S100A4 and liprin beta-1 will be changed using transient transfection and siRNA inhibition. The effect of these perturbations on the overall migration will be monitored in transwell assays. In parallel, FRAP will be carried out using confocal and total internal reflection fluorescence microscopy. The latter will also be used to develop sensitive in vitro binding assays following cell lysis and immobilisation of specific components. Attempts will be made to force cells to take on well-defined shapes by means of patterning the slide surface with substrates such as collagen or fibronectin. Patterns will be made using elastomer stamps created on templates that have been prepared by ion beam focusing. The structural studies should identify specific residues in S100A4 that contribute to binding to specific targets. Mutations will be made to modulate the extent or kinetics of binding and the influence of the same mutations will be determined within cells. While preliminary studies have demonstrated that A431 cells provide a tractable model for analysis using the techniques above, the conclusions drawn will be tested in other cell lines