A novel method to quantitatively measure the binding of proteins to single-stranded DNA (ssDNA) aptamers that employs the inhibition of the DNAzyme hydrolysis of aptamer monolayers is described. A 28-base DNAzyme was designed to specifically bind to and cleave a 29-base ssDNA sequence that can fold into a G-quartet aptamer and bind the protein thrombin. The binding strength of the DNAzyme to the aptamer sequence was designed to be less than the binding strength of the thrombin to the aptamer (Delta G degrees = -43.1 and -51.8 kJ/mol, respectively). Formation of the thrombin aptamer complex was found to block DNAzyme cleavage activity both in solution and in an ssDNA aptamer monolayer. We denote this method for detecting protein aptamer complexation as "DNAzyme footprinting" in analogy to the process of DNase footprinting for the detection of protein-DNA interactions. By attaching a 40-base reporter sequence to the ssDNA aptamer monolayer, the detection of any protein-aptamer complexes remaining on the surface after DNAzyme activity can be greatly enhanced (down to one thrombin-aptamer complex per 10 000 ssDNA molecules corresponding to 100 fM thrombin in solution) by a subsequent surface RNA transcription amplification reaction followed by RNA detection with nanoparticle-enhanced SPR imaging. In addition to RNA transcription, DNAzyme footprinting can be coupled to a wide variety of other nucleic acid surface amplification schemes and thus is a powerful new route for the enzymatically amplified detection of proteins via protein aptamer complex formation.