ABSTRACT
Expressed protein libraries are becoming a critical tool for new target discovery in the pharmaceutical industry. In order to get the most meaningful and comprehensive results from protein library screens, it is essential to have library proteins in their native conformation with proper post-translation modifications. This goal is achieved by expressing untagged human proteins in a human cell background. We optimized the transfection and cell culture conditions to maximize protein expression in a 96-well format so that the expression levels were comparable with the levels observed in shake flasks. For detection purposes, we engineered a 'tag after stop codon' system. Depending on the expression conditions, it was possible to express either native or tagged proteins from the same expression vector set. We created a human secretion protein library of 1432 candidates and a small plasma membrane protein set of about 500 candidates. Utilizing the optimized expression conditions, we expressed and analyzed both libraries by SDS-PAGE gel electrophoresis and Western blotting. Two thirds of secreted proteins could be detected by Western-blot analyses; almost half of them were visible on Coomassie stained gels. In this paper, we describe protein expression libraries that can be easily produced in mammalian expression systems in a 96-well format, with one protein expressed per well. The libraries and methods described allow for the development of robust, high-throughput functional screens designed to assay for protein specific functions associated with a relevant disease-specific activity.
