To efficiently engineer novel proteins, we need fast, scalable, and user-friendly in vitro methods for producing, purifying, and functionally characterizing many protein sequences in parallel. Such methods would be able to make toxic proteins that cannot be screened in cells, enable assays with cell-incompatible conditions (such as denaturants, non-neutral pH, or elevated temperatures), avoid background noise from endogenous proteins, and separate protein folding from function to enable quantitative measurement of protein function. Here, we present a novel bead-based platform—amplicon/protein bead (APB)-display—that enables high-throughput expression, purification, and quantitative in vitro binding measurements for more than 100,000 variants within 3 days. APB-display leverages templated emulsification to rapidly generate hydrogel beads that are each coated with many identical DNA amplicons and immobilized protein molecules corresponding to a single library variant. This bead-displayed protein library can subsequently be sorted and sequenced to enable a wide variety of biochemical and biophysical functional assays (e.g., quantifying folding stability, ligand binding, or catalysis). Here, we optimize and demonstrate the power of APB-display by expressing a protein library with 144,000 FLAG epitope variants as APBs, sorting based on anti-FLAG antibody binding with FACS, and recovering quantitative binding affinities for more than 100,000 variants. Our results agree well with previous studies of FLAG epitope variants, and our method reliably quantifies equilibrium dissociation constants ( K D s) ranging from 1 nM–10 μM. By enabling the functional screening of pooled protein libraries in one-pot reactions with standard equipment and less than a mL of reagents, we anticipate that APB-display will offer time, equipment, and >100-fold cost savings over plate-based methods for ultra-high throughput in vitro protein characterization.
Passow et al. (Sun,) studied this question.