Optimized Yeast Surface Display Workflow for Directed Evolution of Proteases

Podsumowanie

This protocol provides an optimized workflow for the directed evolution of viral proteases in Saccharomyces cerevisiae using a surface-display-based selection system. It outlines DNA insert library and substrate cassette design, electroporation optimized for high transformation efficiency, fluorescence-activated cell sorting-based selection for protease cleavage, and sequencing of enriched variants.

Streszczenie

Directed evolution mimics the natural evolutionary process in a fast, controlled laboratory environment to evolve proteins with desirable functions or traits. Screening a large pool of mutant protein-coding genes allows functional selection of desired variants for applications in biotechnology, medicine, and synthetic biology. Using this protocol, we perform directed evolution in Saccharomyces cerevisiae to engineer protease variants with high specificity for a non-native substrate sequence. We display these protease variants on the yeast cell surface to facilitate the selection of desired variants via fluorescence-activated cell sorting (FACS) over multiple rounds of enrichment. Four key elements of the substrate cassette are co-expressed with the protease library: a fusion protein composed of (i) the yeast adhesion receptor subunit Aga2, (ii) selection and (iii) counter-selection substrate sequences, (iv) epitope tag sequences, and an optional endoplasmic reticulum (ER) retrieval signal sequence. Yeast cells in which only the selection substrate sequence is cleaved are isolated using multicolor FACS via fluorescent (phycoerythrin/fluorescein isothiocyanate–tagged) anti-epitope antibodies. Here we provide details of the optimized, step-by-step protocol to conduct a protease evolution campaign, including DNA insert library design and substrate cassette design, yeast cell electroporation with high transformation efficiency (up to 10⁹ transformants per microgram of DNA), FACS-based selection, enrichment, and sequencing of evolved variants. We demonstrate our directed evolution protocol by evolving the tobacco etch virus nuclear inclusion A protease (TEVp) from cleaving its natural substrate sequence, ENLYFQ↓S, to cleaving a new sequence, ENLYFE↓S.

Wprowadzenie

Cysteine proteases of the Potyviridae family, specifically the nuclear inclusion A (NIa) proteases, represent a largely untapped source of sequence-specific endopeptidases. With more than 3,800 characterized family members, NIa proteases cleave a distinct seven-amino-acid substrate sequence. The potyviral family offers a diverse set of proteases for engineering programmable protein control systems, constructing orthogonal protease-based genetic circuits, and developing sequence-specific biotechnological tools^1,2. Despite this natural diversity, only two NIa proteases, TEVp and tobacco vein mottling virus protease (TVMVp), have been thoroughly characterized and engineered for broad laboratory and biotechnological use^3,4. This limited protease toolkit limits the design space for multicomponent circuits that require orthogonal protease activities⁵, thereby motivating efforts to further expand and characterize NIa proteases through directed evolution^6,7.

Yeast surface display has emerged as one of the most powerful platforms for directed evolution, enabling high-throughput protein library screening via fluorescence-activated cell sorting (FACS)^6,7. For example, the yeast endoplasmic reticulum sequestration screening (YESS) system couples protease activity to surface display through the secretory pathway of S. cerevisiae^6,7. The YESS system has been applied to engineer novel protease-substrate specificities and has demonstrated utility for multi-round directed evolution campaigns. In this system, protease variants are co-expressed with a fusion protein containing a cleavage substrate cassette with epitope markers, and the Aga1p surface anchor is expressed in the EBY100 yeast strain. Differential labeling with fluorescent antibodies distinguishes cleaved from non-cleaved substrate cassettes displayed on the cell surface. This activity-dependent display mechanism is directly compatible with iterative FACS-based selection for libraries of 10⁷ or more variants^6,7.

A fundamental prerequisite for directed evolution is the ability to introduce variant libraries at a sufficient scale to achieve adequate coverage of the targeted sequence space. For site-saturation mutagenesis libraries spanning multiple amino acid positions, transformation yields of 10⁸ to 10⁹ unique transformants are needed to ensure that the diversity introduced into the library is well represented in the sorted population. Several methods exist for introducing exogenous DNA into S. cerevisiae, each with distinct throughput ceilings. The conventional chemical transformation method uses lithium acetate/single-stranded carrier DNA/polyethylene glycol (LiAc/ssDNA/PEG) due to its simplicity and accessibility. However, it typically achieves transformation efficiencies of only 10⁴–10⁶ colony-forming units (CFU) per microgram of DNA, falling short of the sequence variation needed for large library campaigns⁸. By contrast, the electroporation method applies a high-voltage electric field to transiently permeabilize the cell membrane and facilitate DNA uptake. Electroporation consistently achieves the highest transformation efficiencies among available transformation methods, with optimized protocols reporting yields of 10⁸–10⁹ CFU⁹. It has been observed that electroporation efficiency in S. cerevisiae is critically sensitive to cell growth phase, DNA mass, and buffer composition, providing a framework that has since been widely adapted for library-scale directed evolution⁹.

The protocol described here uses S. cerevisiae strain EBY100, a widely adopted host strain for yeast surface display. EBY100 was engineered by Boder and Wittrup by integrating an AGA1 expression cassette under the control of the galactose-inducible GAL1 promoter into the chromosomal AGA1 locus of the protease-deficient parent strain BJ5465¹⁰. The strain is commercially available from the ATCC (catalog no. MYA-4941; depositor K.D. Wittrup), carrying the genotype MATa AGA1::GAL1-AGA1::URA3 ura3-52 trp1 leu2-Δ1 his3-Δ200 pep4::HIS3 prb1Δ1.6R can1 GAL (URA+, leu−, trp−). As a result, both Aga1p and the Aga2p-substrate cassette fusion are expressed from the same galactose-inducible promoter, ensuring that the two subunits are co-induced and assemble into the disulfide-linked Aga1p–Aga2p complex that anchors the displayed fusion to the cell wall.

Despite the widespread use of EBY100 in yeast display applications, no published protocol has been systematically optimized for electroporating this strain at library-scale yields. EBY100-specific parameters, such as the optimal growth phase at harvest, the medium composition during subculture, and the quantitative relationship between initial optical density at 600 nm (OD₆₀₀) and transformation efficiency, remain to be rigorously characterized and optimized. Here, we report that transformation efficiency in EBY100 increases monotonically with the culture's OD₆₀₀ at harvest, with cells in late log phase yielding substantially higher transformation rates than those harvested at early log phase. We evaluated critical protocol parameters, including initial OD₆₀₀ values at inoculation, the use of 2 × Yeast Extract Peptone Dextrose (YPD) medium for subcultures, electroporation buffer formulations, and post-electroporation outgrowth conditions. The protocol presented here provides an optimized, step-by-step workflow (Figure 1A–C, Figure 2, Figure 3A–D, Figure 4A–E) for preparing electrocompetent EBY100 cells and performing high-efficiency DNA library electroporation, yielding transformation rates consistently above 10⁹ CFU. To validate this protocol, we conducted a comprehensive directed evolution campaign to engineer TEVp with altered cleavage specificity across four rounds of FACS-based selection and enrichment. Sequencing of enriched protease variants confirmed substantial diversification at targeted residues within the protease's substrate-binding pocket, demonstrating that the protocol provides sufficient library coverage of sequence space to support productive evolution campaigns. This protocol will be of broad utility to researchers using EBY100 for yeast surface display-based directed evolution and FACS-compatible, high-throughput screening of proteases, antibodies⁹, kinases¹¹, and receptors¹², among other proteins.

Protokół

All experiments involving recombinant S. cerevisiae were conducted in accordance with the policies and procedures of the Institutional Biosafety and Chemical Safety Committee at The University of Texas at Dallas. Recombinant yeast strains were handled under Biosafety Level 1 (BSL-1) containment in accordance with established institutional microbiological practices. Electroporation, culture, and fluorescence-activated cell sorting (FACS) were performed in accordance with approved institutional biosafety procedures. Yeast cultures and contaminated consumables were decontaminated and disposed of as biohazardous waste in accordance with institutional guidelines.

1. Vector and insert preparation

1. Generation of potyviral protease libraries via site-saturation mutagenesis
   1. Design using sequence-design software overlapping 30- to 50-mer, forward and reverse oligonucleotides to generate a mutant library of proteases. Replace the wild-type codon in the corresponding oligonucleotide with an NNS degenerate codon, where N represents A, T, G, or C, and S represents G or C, for positions targeted for mutagenesis.
   2. Design the 3′ overlap between each forward and reverse oligonucleotide pair with a melting temperature (Tm) of 55–60 °C. Design the first and last oligonucleotides to contain a 50-nt overlap with the yeast surface display vector beyond the restriction sites.
   3. Ensure that at least one end of the insert exactly overlaps one end of the linearized vector to facilitate homologous recombination¹³. Avoid gaps between adjacent oligonucleotides¹⁴.
   4. Resuspend oligos to 100 µM with deionized H₂O. In a sterile 1.5 mL tube, combine 0.5 µL of each oligo to a final volume of 200 µL (final concentration of 250 nM for each oligo).
   5. Set up a PCR reaction with the components listed in Table 1 to assemble the protease library, using the thermocycling conditions in Table 2.
   6. Set up a second PCR reaction using the outermost forward and reverse oligos (Table 3) using the first assembled PCR product as a template. Amplify the product as shown in Table 4.
   7. Run the second PCR product on a 1% agarose gel with ethidium bromide (0.5 µg/mL) at 240 V for 30 min. Excise the band corresponding to the expected DNA length.
      NOTE: CAUTION: Ethidium bromide is a mutagen and should be handled using appropriate personal protective equipment (PPE), including a laboratory coat, nitrile gloves, and safety glasses. Avoid skin contact and inhalation. Dispose of ethidium bromide-containing gels, buffers, and consumables as hazardous chemical waste in accordance with institutional safety guidelines.
   8. Purify using a gel extraction kit. Elute in 50 µL of deionized H₂O.
   9. Measure DNA concentration using a spectrophotometer. Store the DNA at −20 °C.
      NOTE: Each 50 µL reaction of the 2^nd PCR would yield about 0.5–1.0 µg of total DNA. Since each electroporation requires 12 µg of the protease library (i.e., insert), scale up reactions accordingly.
2. Substrate cassette assembly
   1. Prepare LB-agar plates and LB-chloramphenicol liquid media. See recipes in Table 5.
   2. Design a set of overlapping 30-mer oligonucleotides in the forward and reverse direction for substrate cassette assembly. Design the 5′ ends of each reverse oligonucleotide and forward oligonucleotide of the next fragment to match by 4 nucleotides.
   3. Resuspend lyophilized oligos to 100 µM using deionized H₂O.
   4. Mix reagents as in Table 6.
   5. Use the cycling program in Table 7 to anneal and phosphorylate the oligo mix.
   6. Dilute annealed oligos 4-fold in deionized H₂O to reach a final concentration of 1.25 µM.
   7. Combine the annealed oligo mix with the vector (e.g., pDD1523), following Table 8.
   8. Perform Golden Gate assembly, following the cycling program in Table 9.
      NOTE: For example, the pDD1523 plasmid has a type IIS restriction site (BsmBI) downstream of the GAL1/10 promoter. Successful assembly allows for galactose induction of the substrate cassette⁷.
   9. Transform the resulting plasmid into E. coli.
   10. Plate cells on LB agar with the corresponding antibiotic. Incubate for 16–18 h at 37 °C.
   11. Pick isolated colonies and inoculate them in 5 mL of LB media. Incubate overnight (16–18 h) at 37 °C with shaking (220 rpm).
   12. Extract plasmids from overnight cultures using a plasmid miniprep kit.
   13. Measure the plasmid concentration using a spectrophotometer, then send for sequencing.
3. Linearization of yeast display vector
   1. Linearize the yeast display plasmid, e.g., digest pDD1524 with SphI and PstI restriction enzymes to prepare the yeast display vector containing the substrate cassette for electroporation. Mix the reagents listed in Table 10 and incubate at 37 °C for 1 h.
   2. Run the restriction digest product on a 1% agarose gel at 240 V for 30 min. Excise the band corresponding to the expected DNA length and purify using a gel extraction kit.
      NOTE: Each 50 µL of restriction digest would yield about 0.5–1.0 µg of total DNA. Since each electroporation requires 4 µg of linearized vector, scale up the digest reactions accordingly.

2. Electroporation

Note: Follow institutional safety procedures for handling biological materials and reagents, wear appropriate PPE, and dispose of biological and chemical waste in accordance with institutional guidelines.

1. Preparation of buffers, plates, and media (2 days prior to electroporation)
   1. Prepare growth media (1 × YPD; 2 × YPD; Synthetic Dextrose Casamino Acids, SDCAA+; and Synthetic Galactose Casamino Acids, SGCAA+), plates (YPD and SDCAA), electroporation buffer, and sterile water following the recipes in Table 5. Store at 4 °C.
2. Preparation of a streaked plate of EBY100.
   1. Streak out EBY100 yeast onto YPD agar from a glycerol stock. Grow at 30 °C for 2 days.
   2. Store the plate at 4 °C after growth.
      NOTE: A streaked plate remains usable for up to 1 month; mark the streak date on the plate.
3. Preparation of EBY100 pre-culture (1 day prior to electroporation)
   1. Prepare 125 mL of 1 × YPD in a 500 mL Erlenmeyer flask.
   2. Prepare 25 mL of 1 × YPD in a 50 mL conical tube. Using a micropipette tip, collect EBY100 cells from the plate grown for 2 days. Then, transfer the yeast cells from the pipette tip into the conical tube containing 1 × YPD.
      NOTE: The streak of cells should form a smear approximately 1–2 mm in length.
   3. Remove the micropipette tip and vortex the conical tube to ensure adequate mixing.
   4. Prepare a 150 mL pre-culture by transferring 25 mL of 1 × YPD containing the yeast cells into the 500 mL flask containing 125 mL of 1 × YPD. Swirl to mix.
   5. Using a spectrophotometer, measure the OD₆₀₀ by preparing two microcuvettes with a path length of 1 cm: one with 1 mL of 1 × YPD as a blank control and another with 1 mL of the pre-culture
   6. Grow 150 mL of pre-culture overnight (16–20 h) at 30 °C with orbital shaking at 270 rpm.
4. Preparation of subculture at desired initial OD₆₀₀ (on the day of electroporation)
   1. Prepare the conditioning buffer and outgrowth media, as described in Table 5.
   2. Measure the OD₆₀₀ of the overnight pre-culture (expected OD₆₀₀ is 17–22). To be within the linear range of the spectrophotometer, measure a 1:20 dilution of the pre-culture.
   3. Calculate the volume of pre-culture required to inoculate a 100 mL subculture. An initial subculture OD₆₀₀ of 3.0–4.0 is recommended.
   4. Dilute the culture to the desired initial OD₆₀₀ in 100 mL of 2 × YPD in a 500 mL Erlenmeyer flask.
   5. Incubate the 100 mL subculture at 30 °C with orbital shaking at 270 rpm for 4 h.
5. Preparation of media during 4 h incubation
   1. Keep ice-cold electroporation buffer and sterile water ready (from step 2.1.1).
   2. Prepare conditioning buffer (100 mL), following recipe in Table 5. Use 20 mL of conditioning buffer per transformation; scale up accordingly. Store at room temperature (20–23 °C).
      NOTE: CAUTION: Conditioning buffer contains dithiothreitol (DTT) and lithium acetate (LiAc), which are chemical irritants that may cause skin and eye irritation. Wear appropriate PPE, including a laboratory coat, nitrile gloves, and safety glasses, and handle these reagents in accordance with institutional chemical safety procedures. Dispose of unused solutions and spent buffers as chemical waste in accordance with institutional guidelines.
   3. Prepare outgrowth media (50 mL): Mix 25 mL of 2 × YPD and 25 mL of 1 M sorbitol together. Use 8 mL of outgrowth media per transformation. Store at room temperature (20–23 °C).
6. Preparation of electrocompetent yeast cells after 4 h incubation.
   1. Split 100 mL subculture into two 50 mL conical tubes. Collect cells by centrifugation at 4 °C and 1,500 × g for 3 min, then decant the medium. Keep cells on ice until step 2.6.4 is completed.
   2. Gently resuspend each cell pellet with 25 mL of ice-cold sterile water. Centrifuge at 4 °C and 1,500 × g for 3 min, then decant the media.
   3. Repeat step 2.6.2.
   4. Gently resuspend each cell pellet with 25 mL of ice-cold electroporation buffer. Centrifuge at 4 °C and 1,500 × g for 3 min, then decant the media.
   5. Remove the cells from ice and gently resuspend each cell pellet with 10 mL of room temperature (20–23 °C) conditioning buffer. Then, pool into a single 125 mL Erlenmeyer flask and incubate at 30 °C with orbital shaking at 270 rpm for 30 min.
   6. Pour the cells into a 50 mL tube. Centrifuge at 4 °C and 1,500 × g for 3 min. Decant the media.
   7. Gently resuspend the cell pellet with 50 mL of ice-cold electroporation buffer. Centrifuge at 4 °C and 1,500 × g for 3 min, then decant the media.
   8. Resuspend the cell pellet to a final volume of 1 mL by adding approximately 100–200 µL of electroporation buffer. Keep cells on ice until electroporation.
      NOTE: Electrocompetent cells can be frozen (i.e., placed in a –80 °C freezer) and used later without significant loss of electroporation efficiency.
7. Electroporation, outgrowth, and plating
   1. Pre-chill electroporation cuvette(s) with a gap size of 0.2 cm on ice.
   2. Transfer 380 µL of the resuspended pellet to a sterile 1.5 mL tube. Mix with 4 µg of plasmid DNA or with 4 µg of linearized plasmid vector and 12 µg of insert. Gently mix by pipetting.
      NOTE: To maximize the number of electroporated cells, vector and insert DNA concentrations should ideally be 700–800 ng/µL. If concentrations are less than that, use a vacuum concentrator.
   3. Transfer suspension into the pre-chilled electroporation cuvette(s). Keep on ice for 5 min.
   4. Wipe the sides of the cuvette after 5 min with a lint-free laboratory wipe to get rid of residual moisture, which may lead to arcing during electroporation. Avoid any bubbles or cells above the metal plates in the cuvette, as they may also lead to arcing. To maintain the cuvette's cold temperature, do not touch the plate electrodes.
   5. Prepare 8 mL of outgrowth media in a 50 mL conical tube.
   6. Electroporate the cells with the following settings: A voltage of 2.5 kV, capacitance of 25 µF, and resistance of 200 Ω⁹. The acceptable time constant range is 2.8–4.5 ms.
      NOTE: CAUTION: Electroporation involves high-voltage electrical pulses. Ensure cuvettes are properly seated and follow the manufacturer's operating instructions. Dispose of used electroporation cuvettes as biohazardous waste according to institutional guidelines.
   7. Flood the cuvette immediately with approximately 200 µL of outgrowth media from the conical tube and transfer electroporated cells into the 50 mL conical tube. Add another 200–350 µL of outgrowth medium until no visible liquid remains in the cuvette.
   8. Incubate the 50 mL conical tube of electroporated cells in 8 mL of outgrowth media at 30 °C without shaking for 1 h. Invert the tube every 15 min to mix.
   9. Collect cells by centrifugation at 1,500 × g, 25 °C for 3 min. Then decant the media.
   10. Resuspend cells in appropriate yeast dropout media to maintain selection for transformed cells. For plasmids containing the TRP1 selection marker, resuspend cells in 250 mL of SDCAA+ (which lacks tryptophan) to selectively recover yeast that retained the plasmid.
   11. Incubate overnight (16–20 h) at 30 °C with orbital shaking at 270 rpm.

3. Induction, staining, and flow cytometric sorting of yeast cells

1. Induction of protease library and substrate cassette (1 day before sorting)
   1. Measure OD₆₀₀ as in step 2.4.2 after overnight growth of transformants in dropout media.
   2. Aliquot 12 mL of yeast transformants and centrifuge at 3,200 × g for 5 min.
   3. Remove the supernatant and resuspend the cell pellet in SGCAA+ induction medium to an OD₆₀₀ of 0.5. Calculate the required resuspension volume using the dilution equation, C₁V₁ = C₂V₂.
   4. Transfer to a 125 mL Erlenmeyer flask and grow overnight for 18–24 h at 30 °C with orbital shaking at 270 rpm.
2. Fluorescent antibody staining (on the day of sorting)
   1. Measure OD₆₀₀ of cells post-induction. Use two cuvettes: one with 1 mL of SGCAA+, and the other with 950 µL of SGCAA+ mixed with 50 µL of cells. Multiply the measured OD₆₀₀ value by 20 to get the true OD₆₀₀ of post-induction cells.
   2. Normalize each sample to 3 × 10⁷ cells for antibody staining and FACS sorting. Use C₁V₁ = C₂V₂ to calculate the volume of cells to transfer from the induction flask.
      NOTE: For four-site NNS saturation mutagenesis, the theoretical library size is 10⁶ variants. To sufficiently sample the library, screen at least 10 times the library size, or 10⁷ cells. Example: if desired C₁V₁ = 3 × 10⁷ cells and C₂ = post-induction OD₆₀₀ of 6 (i.e., 6 × 10⁷ cells/mL), then aliquot 500 µL of induced cells (i.e., V₂ = 3 × 10⁷ cells / 6 × 10⁷ cells/mL = 500 µL).
   3. Transfer calculated volume of cells to a 1.5 mL tube and centrifuge at 5,000 × g for 5 min.
   4. Remove supernatant and resuspend in 500 µL of phosphate-buffered saline (PBS) + 0.5% bovine serum albumin (BSA).
   5. Centrifuge at 5,000 × g for 5 min and repeat step 3.2.4.
   6. Centrifuge at 5,000 × g for 5 min, and in the meantime, complete step 3.2.7.
   7. Prepare an antibody stain mixture in a 1.5 mL tube containing anti-HA fluorescein isothiocyanate (FITC) (0.5 µg/10⁷ cells), anti-FLAG phycoerythrin (PE) (0.05 µg/10⁷ cells), and PBS + 0.5% BSA (100 µL/10⁷ cells). Keep the tube on ice. 
   8. Remove supernatant and resuspend with the stain mixture. Use 300 µL of the stain mixture per 3 × 10⁷ cells. Do not vortex the tube; only invert or flick.
   9. Keep cells on ice in the dark for 1 h after resuspension.
3. Post-staining wash
   1. Centrifuge stained cells at 5,000 × g for 5 min. Remove the supernatant and resuspend the pellet in PBS containing 0.5% BSA using 0.5 mL per 1 × 10⁷ cells (e.g., 500 µL for 1 × 10⁷ cells or 1.5 mL for 3 × 10⁷ cells).
   2. Centrifuge at 5,000 × g, 5 min. Remove supernatant, resuspend in 750 µL of PBS + 0.5% BSA.
   3. Transfer another 750 µL of PBS + 0.5% BSA to a 10 mL collection tube.
4. FACS
   1. Perform necessary startup and preparation of the cytometer for a new experiment.
   2. Create three dot-plots. On the first dot-plot, select the X-axis name to display the forward scatter-area (FSC-A) channel and the Y-axis name to display the side scatter-area (SSC-A) channel.
      NOTE: This will facilitate the selection of live cells.
   3. Keep the X-axis name to display FSC-A channel while changing the Y-axis channel to display forward scatter-height (FSC-H) on the second dot-plot.
      NOTE: This will facilitate selecting singlets.
   4. Select the X-axis name to display the PE channel and the Y-axis name to display the FITC channel on the third dot-plot.
   5. Run negative control on the cytometer using the 530/30 nm (FITC) and 585/42 nm (PE) channels. Perform sorting using a 70-µm nozzle at 70 psi sheath pressure. Acquire samples at a speed of 4000–5000 events/s.
   6. Draw a polygon gate around the main population of intact yeast cells to exclude debris and small particles with low FSC-A and SSC-A signals on the first dot-plot (FSC-A versus SSC-A).
   7. Gate singlets on an FSC-H versus FSC-A plot within this gate, by selecting the linear population exhibiting a proportional relationship between FSC-H and FSC-A. Exclude events that deviate from the linear distribution and correspond to doublets and cell aggregates.
      NOTE: The third dot-plot for PE versus FITC will display a diagonal cluster of dots for the negative control (i.e., yeast cells expressing the intact substrate cassette).
   8. Run samples and sort yeast cells based on desired PE/FITC values into a collection tube(s). Analyze a total number of cells corresponding to at least 10-fold the theoretical library size to ensure adequate representation of all variants (e.g., analyze ~1 × 10⁷ cells for a library with a theoretical diversity of ~1 × 10⁶ variants).
   9. Position the sort gate below the PE/FITC diagonal of the negative control population.
      NOTE: More stringent gates improve enrichment of desired variants but reduce cell recovery.
5. Post-FACS
   1. Centrifuge collection tube(s) at 3,200 × g for 5 min. Carefully remove the supernatant, leaving 1 mL, and resuspend in 5 mL of PBS.
      NOTE: Leave 1 mL of supernatant, as the cell pellet at the bottom will not be visible after centrifugation.
   2. Centrifuge cells again at 3,200 × g for 5 min. Remove the supernatant, leaving 1 mL. Resuspend with 4 mL of SDCAA+.
   3. In a 250 mL Erlenmeyer flask, add 15 mL of SDCAA+. Transfer the resuspended cells from step 3.5.2 into this flask.
   4. Incubate overnight for 18–24 h at 30 °C with orbital shaking at 270 rpm.
   5. Repeat steps 3.1, 3.2, 3.3, and 3.4 for each round of directed evolution.

4. Plasmid recovery and sequencing of individual clones

1. Grow individual colonies
   1. Make a glycerol stock by mixing 500 µL of the liquid culture after step 3.5.4 and 500 µL of sterile 50% glycerol, resulting in a 25% glycerol stock solution. Vortex to thoroughly mix the cells.
   2. Store in the –80 °C freezer overnight. For long-term storage, transfer into liquid nitrogen.
   3. Scrape the glycerol stock using a sterile 1000 µL tip and mix it into 100 µL of SDCAA+. Plate this mixture onto an SDCAA plate and incubate it upside down at 30 °C.
   4. Pick colonies after 3 days, and grow them overnight in 1.5 mL of SDCAA+ in 14 mL tubes.
2. Perform yeast miniprep to extract plasmids
   1. Pellet the overnight culture by centrifuging at 16,000 × g for 1 min and resuspend in 200 µL of Solution 1 from a yeast plasmid purification kit.
   2. Add 5 µL of lytic enzyme solution (5 U/µL).
   3. Incubate in a 37 °C incubator for 4 h on an end-to-end mixer.
   4. Perform a freeze-thaw cycle by freezing at -80 °C for 20 min and thawing at 42 °C for 10 min in a water bath.
   5. Add 200 µL of Solution 2 from a yeast plasmid purification kit, mix by inverting 2–3 times, and let sit for 5 min.
   6. Add 400 µL of Solution 3 from a yeast plasmid purification kit and mix by inverting 2–3 times. Centrifuge at 16,000 × g for 5 min.
   7. Purify plasmid DNA using a purification kit with a high DNA-binding capacity column.
      NOTE: Because of the large amount of sheared genomic yeast DNA coming from the lysed cells, a silica-based column with a higher DNA-binding capacity (35 µg) needs to be used instead of a lower-capacity yeast plasmid DNA purification column (5 µg)¹⁵.
   8. Measure the DNA concentration using a spectrophotometer. DNA yields are 10–30 ng/µL.
3. Transform purified plasmids into chemically competent E. coli cells
   1. Prepare SOC media, following the recipe in Table 5, and LB-chloramphenicol plates. Also, preheat a water bath to 42 °C.
   2. Warm up an LB-chloramphenicol plate for each plasmid at 37 °C for 20 min.
   3. Take out a tube of chemically competent cells from the –80 °C freezer and thaw on ice.
   4. Aliquot 30 µL of competent cells for each plasmid into a 1.5 mL tube on ice.
   5. Pipette 3 µL of plasmid into competent cells. Flick to mix and let it sit for 5 min on ice.
   6. Perform heat shock by briefly incubating the cells in the water bath at 42 °C for 40 s.
   7. Incubate on ice for 2 min.
   8. Add 300 µL of SOC media to the cells and incubate for 15 min at 37 °C at 220 rpm.
   9. Plate 50–100 µL of transformed competent cells onto prewarmed plates.
   10. Incubate the plates at 37 °C upside down for 18 h.
4. Colony picking
   1. Add 5 mL of liquid LB-chloramphenicol media to 14 mL round-bottom culture tubes.
   2. Pick a colony using a sterile 1000 µL tip, and drop it into a culture tube.
   3. Place tubes in a shaker (220 rpm, 30°–45° angle to maximize aeration) at 37 °C for 18 h.
5. Plasmid extraction from bacterial liquid culture
   1. Centrifuge the liquid cultures at 3,200 × g for 4 min. Decant the supernatant.
   2. Extract plasmids from overnight cultures using a plasmid purification kit.
   3. Measure concentration of purified plasmids using a spectrophotometer.
6. Submit plasmids for sequencing
   NOTE: Use services such as whole-plasmid nanopore sequencing, or, alternatively, Sanger sequencing with a primer flanking the mutagenized region.

Wyniki

Efficient library construction is critical to ensure sufficient coverage of the mutational space. To avoid loss of rare functional variants, the number of transformants should reach at least 10× to 100× the theoretical library size. Because transformation efficiency in S. cerevisiae is influenced by cell concentration and is highest in early- to mid-log-phase cells¹⁶, we optimized the initial cell density prior to electroporation. Yeast cultures were subcultured at initial OD₆₀₀ values ranging from 1.0–4.0 and incubated for 4 h (n = 2–4 independent biological experiments), after which the final OD₆₀₀ values were mapped onto the S. cerevisiae growth curve to confirm that cells remained within the mid-log growth phase (Figure 5A, Supplementary Figure 1).

Electroporation of these cultures with 4 µg plasmid DNA (n = 2 independent biological experiments) showed that transformation efficiency differed significantly across initial OD₆₀₀ conditions (one-way ANOVA, p = 8.92 × 10⁻⁷). Relative to cultures initiated at OD₆₀₀ = 1.0, all higher starting cell densities produced significantly greater transformation efficiencies (Dunnett's multiple-comparisons test; OD₆₀₀ = 1.5, adjusted p = 0.0020; OD₆₀₀ ≥ 2.0, adjusted p < 0.0001), with the highest transformation efficiency observed at an initial OD₆₀₀ of 4.0. To take advantage of yeast cells' natural ability to perform homologous recombination, we also electroporated 4 µg of the linearized plasmid vector and 12 µg of the insert (i.e., protease library) into yeast cells (n = 2 independent biological experiments). Transformation efficiency differed significantly among the tested initial OD₆₀₀ conditions (one-way ANOVA, p = 0.0023). Relative to cultures initiated at OD₆₀₀ = 1.0, transformation efficiencies were significantly higher at OD₆₀₀ = 3.0 (adjusted p = 0.0018), 3.5 (adjusted p = 0.0062), and 4.0 (adjusted p = 0.0024) (Dunnett's multiple-comparisons test). We observed the highest transformation efficiency at a moderate initial subculture density (OD₆₀₀ = 3.0), indicating an optimal balance between cell competency and recombination efficiency under these conditions (Figure 5B). Compared with plasmid-only transformation, co-electroporation of a linearized vector and protease library insert (n = 2 independent biological replicates) yielded significantly higher transformation efficiencies across the tested initial OD₆₀₀ conditions (two-way ANOVA on log10-transformed transformation efficiencies, p = 9.40 × 10⁻¹²). On average, co-electroporation produced a mean 121-fold increase in transformation efficiency (95% CI, 63–233-fold) relative to plasmid-only transformation (Figure 5B).

During a FACS run of yeast cells containing the mutant protease library and the substrate selection cassette (Figure 6A), cells will either display a truncated cassette or a protease that still cleaves the counter-selection substrate (Q1), express non-functional protease variants (Q2), or express protease variants that successfully cleave the selection substrate only (Q3). Yeast cells expressing non-functional variants appeared as a dominant population distributed along the diagonal in PE versus FITC plots, reflecting co-retention of both signals (Figure 6A, Q2). By contrast, functional variants that cleaved the selection substrate while sparing the native (counter-selection) substrate formed a distinct off-diagonal population characterized by high-PE and medium-to-low FITC signal intensity (Figure 6A, Q3). Applying a stringent gating strategy to isolate this off-diagonal population resulted in progressive enrichment of functional variants across successive rounds of sorting¹⁰. This enrichment is visually observed as an increasingly defined cluster of cells separated from the diagonal cluster (Figure 6B, left to right). If no distinct off-diagonal population is observed, this may indicate low library quality, insufficient expression, or suboptimal staining conditions. As a negative control, Figure 6C,D shows electroporated, induced, unstained yeast cells and the corresponding FITC and PE fluorescence histograms.

Within the enriched off-diagonal population, subpopulations can be further resolved based on FITC intensity. Cells exhibiting a medium FITC signal can be sorted separately from those with a low FITC signal. Sequencing these distinct populations revealed unique mutation profiles (Table 11), indicating that varying cleavage efficiencies are associated with different protease variants.

Figure 1: Vector and Insert Preparation. (A) Generation of potyviral protease libraries via site-saturation mutagenesis. (B) Substrate cassette assembly and integration into yeast display vector (pDD1523). (C) Linearization of yeast display vector (pDD1524) via restriction digestion. Please click here to view a larger version of this figure.

Figure 2: Electroporation of the substrate cassette and the PCR-amplified mutant protease library into competent EBY100 cells. (Dashed box) In vivo circularization of the plasmid containing both protease and substrate cassette. Please click here to view a larger version of this figure.

Figure 3: Induction, staining, and flow cytometric sorting of yeast cells. (A) Induction of protease library and substrate cassette. (B) Fluorescent antibody staining: Staining with anti-HA fluorescein isothiocyanate (FITC) and anti-FLAG phycoerythrin (PE) antibodies. (C) Stained cells are sorted using Fluorescence-Activated Cell Sorting (FACS). (D) Recovery of FACS-sorted cells in selection media. Please click here to view a larger version of this figure.

Figure 4: Plasmid recovery and sequencing of sorted yeast cells. (A) Plating of sorted yeast cells. (B) Plasmid extraction from individual yeast colonies. (C) E. coli transformation of extracted plasmids. (D) Plasmid extraction from individual E. coli colonies. (E) Nanopore sequencing of extracted plasmids. Flask and tube icons adapted from DBCLS and Helicase 11, respectively, via Bioicons (https://bioicons.com) and adapted under the terms of the Creative Commons Attribution 4.0 International License (CC BY 4.0). Please click here to view a larger version of this figure.

Figure 5: Yeast growth curves in different growth media and the influence of initial Optical Density at 600 nm (OD₆₀₀) of subculture on yeast electroporation efficiency. (A) Growth curves of yeast in 1 × Yeast Extract Peptone Dextrose (YPD), 2 × YPD, Synthetic Dextrose Casamino Acids Agar without antibiotics (SDCAA-), and Synthetic Dextrose Casamino Acids Agar with antibiotics (SDCAA+). (B) Transformation efficiency of yeast electroporated with 4 µg plasmid vector at various subculture initial OD₆₀₀, and the transformation efficiency using linearized vector (4 µg) and protease library insert (12 µg) at the same initial OD₆₀₀ values. Data in (A) are presented as OD₆₀₀ ± SD from two independent biological experiments (n = 2). Data in (B) are presented as mean number of transformants per electroporation on a logarithmic y-axis, with individual biological replicates shown as circles (n = 2). Statistical significance for the plasmid and vector-plus-insert electroporation optimization experiments was determined using one-way ANOVA on log10-transformed electroporation efficiencies followed by Dunnett's multiple-comparisons test using OD₆₀₀ = 1.0 as the control. ns, not significant; p < 0.01 (**); p < 0.0001 (****). Statistical significance of the difference between plasmid-only and vector-plus-insert transformation efficiencies was determined using two-way ANOVA on log10-transformed transformation efficiencies. p < 0.0001 (****). Please click here to view a larger version of this figure.

Figure 6: Fluorescence-Activated Cell Sorting (FACS)-based enrichment of yeast clones. (A) Schematic representation of the gating strategy. Cells were divided into three regions: Q1, phycoerythrin (PE)-low cells; Q2, PE-high/fluorescein isothiocyanate (FITC)-high cells; and Q3, PE-high cells with low-to-intermediate FITC fluorescence. Cells within Q3 were collected for enrichment. (B) Representative FACS plots of one independent evolutionary campaign showing sequential rounds of sorting with increasing FITC gating stringency. Mid- and low-FITC cells were collected separately for sequencing. Percentages indicate the fraction of live, single yeast cells within the gated population. (C) Representative FACS plot showing electroporated, unstained yeast cells as a negative control. (D) Histograms of FITC-A (left) and PE-A (right) fluorescence intensities corresponding to the data shown in (C). Please click here to view a larger version of this figure.

Table 1: Reaction mixture for 1st-round PCR amplification of the protease library. Composition of the PCR reaction used for the first-round amplification of the protease library, including the final concentrations and amounts of each reagent in a 50 µL reaction. Please click here to download this Table.

Table 2: Cycling conditions for 1st-round PCR amplification of the protease library. The thermal cycling program was used for the first-round PCR amplification of the protease library. Please click here to download this Table.

Table 3: Reaction mixture for 2nd-round PCR amplification of the protease library. Composition of the PCR reaction used for the second-round amplification of the protease library, including the final concentrations and amounts of each reagent in a 50 µL reaction. Please click here to download this Table.

Table 4: Cycling conditions for 2nd-round PCR amplification of the protease library. The thermal cycling program was used for the second-round PCR amplification of the protease library. Please click here to download this Table.

Table 5: Media and Reagents for Yeast Electroporation and Growth. Tabular description of the mass required from each chemical component to formulate specific amounts of 1 × and 2 × Yeast Extract Peptone Dextrose (YPD), electroporation buffer, conditioning buffer, outgrowth media, Synthetic Dextrose Casamino Acids with antibiotics (SDCAA+), Synthetic Galactose Casamino Acids with antibiotics (SGCAA+), Lysogeny Broth (LB) media, LB-chloramphenicol media, Super Optimal Broth (SOB) media, and Super Optimal Broth with Catabolite repression (SOC) media. Instructions for preparing stock solutions, such as 2 M Sorbitol, 1 M dithiothreitol (DTT), and 2 M lithium acetate (LiAc), are included. Additional instructions for preparing SDCAA, LB, and YPD agar plates are provided. Please click here to download this Table.

Table 6: Reaction mixture for annealing and phosphorylation of oligos. Composition of the reaction mixture used for simultaneous annealing and phosphorylation of oligonucleotides for substrate cassette assembly. Please click here to download this Table.

Table 7: Cycling conditions for annealing and phosphorylation of oligos. Temperature program used for oligonucleotide annealing and phosphorylation for substrate cassette assembly. Please click here to download this Table.

Table 8: Reaction mixture for Golden Gate assembly of substrate cassette and vector plasmid. Composition of the Golden Gate assembly reaction used to ligate the substrate cassette into the vector plasmid. Please click here to download this Table.

Table 9: Cycling conditions for Golden Gate assembly of substrate cassette and vector plasmid. Thermal cycling program used for Golden Gate assembly. Please click here to download this Table.

Table 10: Reaction mixture for linearization of vector plasmid. Composition of the restriction digestion reaction used to linearize the vector plasmid prior to library construction. Please click here to download this Table.

Table 11: Nanopore sequencing results of TEVp variants cleaving ENLYFES. Plasmids were extracted from yeast cells after the fourth and final FACS session. Mutations in each clone are shown relative to the wild-type sequence. Please click here to download this Table.

Supplementary Figure 1: Relationship between initial and final EBY100 culture density during subculture. EBY100 cultures were subcultured at varying initial OD₆₀₀ values, and the final OD₆₀₀ was measured after incubation under identical growth conditions (30 °C, 220 rpm). Final culture density increased linearly with increasing initial OD₆₀₀ (R² = 0.969, p = 5.77 × 10⁻⁵). Data points represent means ± SD of (n = 2–4) independent biological replicates.Please click here to download this file.

Supplementary Figure 2: Rational design of the TEVp mutagenesis library. Cartoon representation of TEVp (PDB ID: 1LVB) with the four residues (gold) <5Å away from the P1 residue (purple) targeted for site-saturation mutagenesis.Please click here to download this file.

Dyskusja

Directed evolution is a high-throughput method that uses iterative cycles of diversification and selection to identify proteins with enhanced or novel biochemical function¹⁷. In this protocol, we detail an optimized workflow to evolve protease-substrate recognition using a cell-surface display system in S. cerevisiae. The surface display system maps protease activity to a detectable fluorescence-based phenotype, enabling FACS-based enrichment of desired clones. We provide the detailed steps for protease library and substrate cassette assembly, high-efficiency electroporation, FACS gating and sorting, and final sequencing of protease variants.

One of the most essential steps in directed evolution is ensuring sufficient library diversity and coverage of potential mutations¹⁷. Our electroporation efficiency experiments show that an initial subculture OD₆₀₀ of 3.0 yields the highest transformation efficiency, exceeding 10⁹ CFU. This scale, achieved with optimized yeast electroporation, substantially exceeds that of conventional chemical transformation methods and enables adequate coverage of large combinatorial libraries^8,9. Accurate FACS gating and sorting are essential for separating cells that display the desired cleavage properties from those that exhibit nonspecific cleavage. During multiple rounds of FACS-based selection, the gating strategy becomes increasingly stringent to selectively enrich for desired protease variants. If gating is too broad, undesired cells may be sorted along with the desired variants. Stringent gating (e.g., increasingly lower thresholds for the FITC signal across rounds) allows the cell population to become enriched for protease variants with higher specificity and activity toward the new target substrate sequence¹⁰. Consistent with past campaigns^7,18, we performed NNS-codon mutagenesis of four residues in the substrate-binding pocket of TEVp that interact with the substrate P1 residue, Q (T146, D148, H167, S170) (Supplementary Figure 2). The FACS plots shown in Figure 6B represent a single representative sorting experiment illustrating the gating strategy and enrichment workflow. Intriguingly, our campaign results produced TEVp variants with a different set of amino acid mutations (Table 11). These results support a key feature of protease evolution: multiple evolutionary paths can lead to the same goal.

Modifications and troubleshooting strategies may be needed when conducting the experiments. For example, during electroporation, working on ice when indicated and carefully resuspending the cell pellets at each step ensures the cells are alive and healthy prior to electroporation. Yeast cells are delicate throughout the electroporation steps until the outgrowth recovery step. Additionally, modification of the FACS sheath fluid may be required to prevent yeast cell flocculation. Sheath fluid is commonly used in FACS machines and may contain preservatives that inhibit bacterial and fungal growth. Therefore, sheath fluid can be replaced with pure PBS to prevent cell death during sorting.

The yeast surface display system requires properly folded proteins to successfully traverse the secretory pathway. Some protease variants may lose activity due to misfolding, leading to false positives during FACS selection caused by nonspecific cleavage or background fluorescence. Well-characterized sources of false positives are worth noting for plasmid-based campaigns like this one. Because cleavage is read out as loss of the C-terminal epitope tag, any mutation that disrupts that tag yields the same low-FITC phenotype as a genuine cleavage event; such tag mutations arise during the large-scale homologous recombination and can enrich across successive sorting rounds. Chromosomal integration of the constant substrate cassette has recently been shown to mitigate this issue¹⁹. The protocol relies on using an insert library while keeping the vector (containing the substrate cassette) constant. Therefore, this protocol is not suitable for those who want to evolve both protease and substrate libraries.

The YESS platform pioneered a general route to reprogram protease-substrate specificity¹⁸. Recent improvements include: adding titratable enzyme/substrate control in YESS2.0⁷, coupling to next-generation sequencing, enabling high-resolution specificity profiling²⁰, and chromosomally integrating the reporters to reduce expression noise and false positives¹⁹. Other orthogonal strategies, such as phage-assisted continuous evolution, have also been applied in engineering protease specificity for therapeutic targets²¹. These tools have found applications beyond TEV protease engineering, such as high-resolution profiling of the SARS-CoV-2 main protease to guide inhibitor design²². A crucial determinant of any such campaign is generating enough transformants to cover large combinatorial libraries (>10⁹), which had not been systematically optimized in the EBY100 S. cerevisiae strain. By defining the growth-phase and electroporation conditions that drive EBY100 transformation above 10⁹ CFU, this protocol removes that bottleneck, enabling comprehensive FACS-based screening of millions of variants.

Materiały

Lista materiałów użytych w tym artykule

Nazwa	Firma	Numer katalogowy	Komentarze
Reagents
10 mM dNTP Mix	NEB	N0447S	PCR reactions (Tables 1, 3)
10X PBS, pH 7.4	Fisher	BP3994	Staining buffer base (Steps 3.2–3.3)
5X Q5 Reaction Buffer	NEB	B9027S	Sold separately; also included with Q5 polymerase
Agar, Bacto	BD Difco	281230-500g	SDCAA agar plates (Table 5); 15 g/L
Agarose (Low-EEO/Multi-Purpose/Molecular Biology Grade)	Fisher BioReagents	BP160-500	Gel Electrophoresis
Anti-FLAG PE Antibody	BioLegend	637309	FACS staining; PE-conjugated anti-FLAG
Anti-HA Tag Antibody [FITC]	GenScript	A01621	FACS staining; FITC-conjugated anti-HA
Bacto Casamino Acids	BD Difco	223120	SDCAA+, SGCAA+, SCAA media (Table 5); 5 g/L
Bacto Peptone / Tryptone	RPI	T60060-10000.0	LB/SOB media preparation; 20 g/L in YPD (Table 5)
BSA, Bovine Serum Albumin, Fraction V	Sigma-Aldrich	126575-10GM	0.5% in PBS; staining buffer (Steps 3.2–3.3)
BsmBI-v2 (200 U)	NEB	R0739S	Golden Gate assembly of substrate cassette (Table 8)
Calcium chloride solution (1M)	Sigma-Aldrich	21115-100mL	Electroporation buffer: 1 mM final (Table 5)
ColiRollers® Plating Beads	Sigma-Aldrich	71013-3	Sterilize by putting in reagent bottles and autoclaving at 121ºC, 15 psi, for 15 mins.
D-Galactose	Thermo Fisher	A12813.30	250 g; 20% w/v stock for SGCAA+ induction (Table 5)
D-Sorbitol	Thermo Fisher	1327300.10	Electroporation buffer (1 M) and outgrowth media (Table 5)
Dextrose (D-Glucose), anhydrous	Sigma-Aldrich	G8270-1KG	YPD, SDCAA+, 20% w/v dextrose stock (Table 5)
Difco Yeast Nitrogen Base w/o Amino Acids	Sigma	Y0626-250G	6.7 g/L in SDCAA/SGCAA (Table 5)
Difco YPD Broth (pre-mixed)	BD	242820-500g	Pre-made YPD alternative; 500 g
DL-Dithiothreitol (DTT)	Sigma-Aldrich	D0632-1G	Conditioning buffer: 10 mM final; aliquot 250 μL, store −20°C
E.Z.N.A. CyclePure PCR Purification Kit	Omega Bio-tek	D6492-02	PCR product cleanup
E.Z.N.A. Gel Extraction Kit	Omega Bio-tek	D2500-01	Gel purification post-PCR and restriction digest (Steps 1.1.8, 1.3.2)
E.Z.N.A. Plasmid DNA Mini Kit I	Omega Bio-tek	D6942-02	Plasmid purification (Step 4.2)
Glycerol (≥99%)	Fisher	BP229-4	50% v/v stock for glycerol stocks (Step 4.1.1)
Kanamycin Sulfate	Fisher	BP906-5	SDCAA+ and SGCAA+: 50 μg/mL final; replaces Sigma K0879-5G
KLD Enzyme Mix	NEB	M0554S	Kinase-Ligase-DpnI; site-directed mutagenesis
KLD Reaction Buffer	NEB	B0554A	Used with KLD Enzyme Mix
LB Broth (Lennox)	Sigma-Aldrich	L3022-1KG	E. coli liquid culture
Lithium Acetate Dihydrate (LiOAc·2H2O)	Thermo Fisher	A17921.30	Conditioning buffer: 0.1 M final; add before DTT (Table 5)
Luria Agar (Luria-Bertani Agar)	RPI	L24020-2000.0	E. coli plating; replaces Sigma LB Agar Lennox L2897-1KG
OmniPur Casamino Acids	Sigma	2240-500GM	Alternative casamino acids source
pDD1523	Zenodo	10.5281/zenodo. 20746804	pCTCon2-TRP-CEN/ARS-Gal10-SPAga2-BsaI; Gal1-Aga2-sfGFP
pDD1524	Zenodo	10.5281/zenodo. 20746804	pCTCon2-Aga2-6xHis-ENLYFQS-FLAG-ENLYFES-HA-ERS
Penicillin-Streptomycin Solution, 100X	Corning	30-002-CI	SDCAA+, SGCAA+: 1X final
PstI-HF (5000 U)	NEB	R3140S	Vector linearization (Table 10); use HF version
Q5 HF DNA Polymerase	NEB	M0492S	High-fidelity polymerase; alternative to Q5 Hot Start
Q5 High GC Enhancer	NEB	B9028A	For high GC-content PCR reactions
Q5 Hot Start DNA Polymerase	NEB	M0491L	PCR assembly and amplification (Tables 1, 3)
rCutSmart Buffer (10X)	NEB	B6004S	Supplied with PstI-HF and SphI-HF (Table 10)
SalI-HF	NEB	R3138L	Additional restriction enzyme
SOC Medium	NEB	B9020S	Post-transformation recovery
Sodium Chloride (NaCl)	Fisher	S271-1	LB / SOB media preparation
Sodium Hydroxide (NaOH)	Fisher	S318-500	Media pH adjustment
Sodium Phosphate Dibasic (Na2HPO4)	Sigma-Aldrich	S9763-500G	SDCAA plates, SCAA media: 38 mM (Table 5)
Sodium Phosphate Monobasic, anhydrous	VWR (BDH)	BDH4542-1KGP	SDCAA plates, SCAA media: 62 mM; replaces NaH2PO4·H2O monohydrate
SphI-HF (1000 U)	NEB	R3182S	Vector linearization in tandem with PstI-HF (Table 10)
Streptomycin	Life Technologies / Sigma	S6501-5g	Antibiotic
T4 DNA Ligase Buffer (10X)	NEB	B0202S	Table 6
T4 Polynucleotide Kinase (500 U)	NEB	M0201S	Table 6
T7 DNA Ligase (3000 U/mL)	NEB	M0318S	Table 8
Yeast Extract	RPI	Y20020-5000.0	YPD (10 g/L) and LB/SOB media
Zymolyase (1000 U)	Zymo Research	E1004-A	Yeast cell wall digestion for plasmid extraction (Step 4.2.2)
Zymoprep Yeast Plasmid Miniprep II	Zymo Research	D2004	Yeast plasmid extraction; includes Solutions 1, 2, 3 (Step 4.2)
ZymoPURE II Plasmid Midi Kit	Zymo Research	D4201-B	Mid-scale plasmid prep; for larger-volume preparations
Equipment and Consumables
Disposable Cuvettes, 1.5 mL	Fisher	14-955-127	For measuring OD600 of liquid cultures
Erlenmeyer Flask, 125 mL	VWR	10536-912	Pre-culture and conditioning incubation
Erlenmeyer Flask, 250 mL	VWR	10536-914	Yeast subculture
Erlenmeyer Flask, 250 mL	VWR	75809-646	Wide-mouth variant
Erlenmeyer Flask, 500 mL	VWR	10536-926	Main pre-culture flask (Step 2.3.1, 2.4.3)
Falcon 50 mL Conical Tubes	Falcon	352070	Cell washes, centrifugation throughout protocol
Flow Cytometry Sorter: BD Fusion	BD Biosciences	FACSAria Fusion Flow Cytometer (catalog number not applicable)	For Fluorescence-Activated Cell Sorting (FACS)
Gene Pulser/MicroPulser Electroporation Cuvettes, 0.2 cm gap	Bio-Rad	1652086	For electroporation of electrocompetent EBY100
GenePulser Xcell Electroporation System	Bio-Rad	1652660	INSTRUMENT; 2.5 kV, 25 μF, 200 Ω (Step 2.7.6)
New Brunswick Innova 40R - Benchtop Orbital Shaker	Eppendorf	M12990084	For yeast liquid culture incubation
Petri Dishes, 100 mm	Fisher	FB0875712	YPD and SDCAA agar plates
SpectraMax QuickDrop UV-Vis Spectrophotometer	Molecular Devices	SpectraMax QuickDrop	For measuring OD600 of liquid cultures and DNA concentration
Vacufuge plus	Eppendorf	2231001204	For concentrating DNA
Software
BD FACSDiva (Version: 8.0)	BD Biosciences	https://www.bdbiosciences.com/en-us/products/software/instrument-software/bd-facsdiva-software	Collection of tools for flow cytometer and application setup, data acquisition, and data analysis
Benchling (Version: Web-based platform, continuous release)	Benchling, Inc.	https://www.benchling.com/	Sequence design software
SnapGene (Version: 8.0.1)	SnapGene, Inc.	https://www.snapgene.com/	Sequence design software