Proteins acting on DNA need to penetrate a dense network of chromatin and associated macromolecules in the cell nucleus to access their target sites. Intracellular mobility of proteins is characterized by diffusion coefficients of the order of 1-100 μm2/s, leading to millisecond time scales for movement on the submicrometer scale. Typical microscopic methods used for characterizing intracellular protein mobility are, e.g., fluorescence photobleaching recovery (FRAP) and fluorescence correlation spectroscopy (FCS). Of these, FRAP can image protein mobility in entire two-dimensional sections of live cells, but is typically limited to the time resolution of confocal image series, some frames per second. FCS, on the other hand, has fast time resolution but so far has been limited to single-point measurements in the focus of a laser beam. Although we have demonstrated first protein mobility maps by point-to-point FCS, this method is extremely time-consuming and not very feasible for live cell measurements. Here we show results from single plane illumination microscopy based fluorescence correlation spectroscopy (SPIM-FCS), a new method for imaging FCS in 3D samples that combines the fast time resolution of FCS with the possibility of acquiring the mobility data in parallel on an entire twodimensional cross-section. This will then provide diffusion coefficients, flow velocities and concentrations in an imaging mode. Extending this technique to two-color fluorescence cross-correlation spectroscopy (SPIM-FCCS) also allows one to measure molecular interactions in an imaging mode