Abstract
Background: The specific binding of proteins to DNA is a key step for many cellular activities, such as transcription regulation, DNA replication, recombination, repair, and restriction. The detection of DNA-binding proteins, as well as the identification of specific binding sites, is therefore important to understand gene expression mechanisms and cellular function. We describe an ultrasensitive method for quantification of DNA-binding proteins.
Methods: We combined the common exonuclease III (ExoIII) footprinting assay and real-time PCR for quantification of DNA-binding proteins, for an assay that does not require antibodies against the target proteins. Double-strand DNA probes were designed to monitor the activities of DNA-binding protein. The protein-binding site is at the 5′ end of the forward primer. When a target protein is present, it will specifically bind to the protein-binding site and produce a physical hindrance to ExoIII, which protects the reverse DNA strand from digestion by ExoIII. The remaining single-strand DNA template can be quantitatively detected by real-time PCR. Conversely, in the absence of the target protein, the naked primer regions will be degraded by ExoIII, which then cannot be amplified by real-time PCR.
Results: We detected the binding of 10 different transcription factors in crude cell extracts. The assay quantitatively detected binding at femtomolar concentrations of protein.
Conclusions: This technique is customizable and easy to establish. It has potential applications in research, medical diagnosis, and drug discovery.
Photoaffinity labeling of transcription factors by DNA-templated crosslinking
