Abstract
Yeast surface display is a promising biotechnological tool that uses genetically modified yeast cell wall proteins as anchors for enzymes of interest, thereby transforming yeast cell wall into a living catalytic material. Here, we present a comprehensive protocol for quantifying surface-displayed β-lactamase on the cell wall of model yeast Saccharomyces cerevisiae. We use β-lactamase as a reporter enzyme, which we tagged to be anchored to the cell wall closer to its N or C terminus, through the portion of the Pir2 or Ccw12 cell wall proteins, respectively. The catalytic activity of surface-displayed β-lactamase is assessed by its ability to hydrolyze nitrocefin, which produces a colorimetric change that is quantitatively measured by spectrophotometric analysis at 482 nm. This system enables precise quantification of the potential of S. cerevisiae strains for surface display, continuous real-time monitoring of enzyme activity, and facilitates the study of enzyme kinetics and interactions with inhibitors within the cell's native environment. In addition, the system provides a platform for high-throughput screening of potential β-lactamase inhibitors and can be adapted for the visualization of other enzymes, making it a versatile tool for drug discovery and bioprocess development.