Forces are increasingly recognized as major regulators of cell structure and function, and the mechanical properties of cells, such as cell stiffness, are essential to the mechanisms by which cells sense forces, transmit them to the cell interior or to other cells, and transduce them into chemical signals that impact a spectrum of cellular responses. Here we reported a new whole-cell cell stiffness measurement technique with a subcellular spatial resolution. This technique was based on a novel cell stretching device that allowed for quantitative control and real-time measurements of mechanical stimuli and cellular biomechanical responses. Our strategy involved a microfabricated array of silicone elastomeric microposts integrated onto a stretchable elastomeric membrane. Using a computer-controlled vacuum, this micropost array membrane (mPAM) was activated to apply equibiaxial cell stretching forces to adherent cells attached on the tops of the microposts. The micropost top positions before and after mPAM stretches were recorded using fluorescence microscopy and further utilized to quantify local cell stretching forces and cell area increments. A robust computation scheme was developed and implemented for subcellular quantifications of cell stiffness using the data of local cell stretching forces and cell area increments generated from mPAM cell stretch assays. Our cell stiffness studies using the mPAM revealed strong positive correlations among cell stiffness, cellular traction force, and cell spread area, and illustrated the important functional roles of actin polymerization and myosin II-mediated cytoskeleton contractility in regulating cell stiffness. Collectively, our work reported a new approach for whole-cell stiffness measurements with a subcellular spatial resolution, which would help likely explain the complex biomechanical functions and force-sensing mechanisms of cells and design better materials for cell and tissue engineering and other applications in vivo.