Measuring the capacity of yeast for surface display of cell wall-anchored protein isoforms by using β-lactamase as a reporter enzyme

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 beta-lactamase on the cell wall of model yeast Saccharomyces cerevisiae. We use beta-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 beta-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 beta-lactamase inhibitors and can be adapted for the visualization of other enzymes, making it a versatile tool for drug discovery and bioprocess development. Yeast surface display (YSD) uses genetically modified yeast to immobilize recombinant enzymes on the cell surface. To quantify the efficiency of YSD, beta-lactamase was fused to selected cell wall proteins and enzyme activity was measured spectrophotometrically at 482 nm using nitrocefin as substrate. This system enables the monitoring of enzyme activity, high-throughput screening of inhibitors, and versatile applications in bioprocess. image