We report the first topographical data of the surface of living endothelial cells at sub-light-microscopic resolution, measurements essential for a detailed understanding of force distribution in the endothelium subjected to flow. Atomic force microscopy was used to observe the surface topography of living endothelial cells in confluent monolayers maintained in static culture or subjected to unidirectional shear stress in laminar flow (12 dyne/cm2 for 24 hours). The surface of polygonal unsheared cells was smooth, with mean excursion of surface undulation between peak height (over the nucleus) and minima (at intercellular junctions) of 3.4±0.7 μm (mean±SD); the mean height to length ratio was 0.11±0.02. In cells that were aligned in the direction of flow after a 24-hour exposure to laminar shear stress, height differentials were significantly reduced (mean, 1.8±0.5 μm), and the mean height to length ratio was 0.045±0.009. Calculation of maximum shear stress and maximum gradient of shear stress (∂τ/ ∂x, where τ is shear stress at the cell surface) at constant flow velocity revealed substantial streamlining of aligned cells that reduced ∂r/∂x by more than 50% at a nominal shear stress of 10 dyne/cm2. Aligned cells exhibited ridges extending in the direction of flow that represented imprints of submembranous F-actin stress-fiber bundles mechanically coupled to the cell membrane. The surface ridges, ≈50 nm in height and 200 to 1000 nm in width, were particularly prominent in the periphery of the aligned cells. These observations (1) represent the first measurement of endothelial surface topography in living cells, (2) demonstrate significant changes in surface topography as a result of exposure to hemodynamic forces, probably as a result of submembranous cytoskeletal reorganization, and (3) facilitate computation of detailed cell-surface force distribution.