Experiments

Protocols

Strain Cultivation

• Escherichia coli

  Materials (all sterilized by autoclaving)

  Table: LB liquid media (for 1 liter)

  Material	Quantity
  yeast extract	5 g
  tryptone	10 g
  NaCl	10 g

  Table: LB agar plate without ampicillin (for 1 liter)

  Material	Quantity
  yeast extract	5 g
  tryptone	10 g
  NaCl	10 g
  agar	15 g

  Table: LB agar plate with ampicillin (for 1 liter)

  Material	Quantity
  yeast extract	5 g
  tryptone	10 g
  NaCl	10 g
  agar	15 g
  1000× ampicillin stock	1 mL

  Procedure

  1. Recovery

  • Thaw Escherichia coli glycerol stocks rapidly in a 37 °C water bath with gentle agitation until just melted.

  • Inoculate 100 µL of the thawed culture into 900 µL of liquid LB medium and incubate at 37 °C with shaking at 200 rpm for 1 hour.

  • Centrifuge the culture at 5000 × g for 1 minute at room temperature.

  • Remove approximately 900–950 µL of the supernatant, leaving 50–100 µL. Resuspend the pellet by gently pipetting.

  • Spread the resuspended cells onto LB agar plates supplemented with ampicillin. Allow the suspension to absorb, then incubate the plates inverted at 37 °C overnight (12–16 h).

  2. Maintenance

  • Spread the revived single colony onto an LB agar plate containing ampicillin, seal the edges with parafilm, and store at 4 °C.

  3. Cultivation

  • Pick a single colony and inoculate it into 5 mL of LB medium containing 100 μg/mL ampicillin. Incubate overnight at 37 °C with shaking at 230 rpm.

  • Inoculate the overnight culture into 100 mL of LB medium containing 100 μg/mL ampicillin at a 1:100 dilution and measure the initial OD₆₀₀.

  • Incubate at 37°C with 200 rpm shaking until the OD₆₀₀ reaches 0.5-0.6.

  4. Preservation

  • Add 20 μL of 100% glycerol to the remaining cells and store at -80 °C.

• Saccharomyces

  Material

  Table: Stock solution (all sterilized by filtration)

  Stock	Composition (for 1 liter)
  40% galactose stock	400.0 g galactose
  20% raffinose stock	200.0 g raffinose
  20% glucose stock	200.0 g glucose
  Amino acid & nucleotide stock	100 mg L-Leucine/20 mg L-Histidine/20 mg L-Methionine/20 mg Uracil

  Table: SC liquid media (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  all 4 amino acid / nucleotide stock	10 mL each
  DO supplement	0.6 g

  Table: SC agar plate (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  all 4 amino acid / nucleotide stock	10 mL each
  DO supplement	0.6 g
  Agar	15 g

  Table: SC-Ura-Leu-Met-His media (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  DO supplement	0.6 g

  Table: SC-Ura media (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  all 3 amino acid stock	10 mL each
  DO supplement	0.6 g

  Table: SC-Leu media (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  Ura, Met, His stock	10 mL each
  DO supplement	0.6 g

  Table: SD-Ura activation media

  Material	Quantity
  SC-Ura liquid media	45 mL
  20% glucose stock	5 mL

  Table: SC-Ura+Raf scale-up media

  Material	Quantity
  SC-Ura liquid media	450 mL
  20% raffinose stock	50 mL

  Table: SD-Leu activation media

  Material	Quantity
  SC-Leu liquid media	45 mL
  20% glucose stock	5 mL

  Table: SC-Leu+Raf scale-up media

  Material	Quantity
  SC-Leu liquid media	450 mL
  20% raffinose stock	50 mL

  Table: SD-Ura agar plate (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  all 3 amino acid stock	10 mL each
  DO supplement	0.6 g
  20% glucose stock	100 mL
  Agar	15 g

  Table: SD-Leu agar plate (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  Ura, Met, His stock	10 mL each
  DO supplement	0.6 g
  20% glucose stock	100 mL
  Agar	15 g

  Table: SC-Ura one-step induction media

  Material	Quantity
  SC-Ura liquid media	180mL
  30% galactose-raffinose mixture (Gal:Raf=2:1)	20mL

  Table: SC-Leu one-step induction media

  Material	Quantity
  SC-Leu liquid media	180mL
  30% galactose-raffinose mixture (Gal:Raf=2:1)	20mL

  Table: SD (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  all 4 amino acid / nucleotide stock	10 mL each
  DO supplement	0.6 g
  20% glucose stock	100 mL
  Agar (for plates only)	15 g

  Table: SD-Ura-His-Leu (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  Met stock	10 mL
  DO supplement	0.6 g
  20% glucose stock	100 mL
  Agar (for plates only)	15 g

  Table: SD-Ura-His (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  Met, Leu stock	10 mL each
  DO supplement	0.6 g
  20% glucose stock	100 mL
  Agar (for plates only)	15 g

  Table: SD-Ura (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium sulfate	5 g
  Met, Leu, His stock	10 mL each
  DO supplement	0.6 g
  20% glucose stock	100 mL
  Agar (for plates only)	15 g

  Table: YPD (for 500 mL)

  Material	Quantity
  BeyoPure™ YPD Broth premixed powder (namely, 5 g/500mL yeast extract, 10 g/500mL peptone, 10 g/500mL D-glucose)	1 bottle
  agar for plates	10 g

  Table: YPD+G418 (for 500 mL)

  Material	Quantity
  BeyoPure™ YPD Broth premixed powder	1 bottle
  G418 stock	2.5 g

  Procedure

  1. Recovery

  • Streak the Saccharomyces cerevisiae glycerol stock stored at -80 °C onto a YPD agar plate, and incubate inverted at 30 °C for 2–4 days to reactivate the strain.

  2. Maintenance

  • Spread the single colony onto the appropriate agar plates, seal the edges with parafilm, and store at 4 °C.

  3. Cultivation

  S. cerevisiae

  • Using a sterile toothpick, pick a single colony and inoculate it into 5 mL of the appropriate plasmid activation medium. Incubate overnight at 30 °C with shaking at 220-240 rpm.

  S. Boulardii

  • Using a sterile toothpick, pick a single colony and inoculate it into 5 mL of liquid YPD medium in a 5 mL culture tube.
  • Mix by vortexing and incubate overnight at 37 °C with shaking at 220-240 rpm.

  4. Preservation

  • Add 50% glycerol to a final concentration of 25% (mix at a 1:1 ratio) and mix thoroughly, then store at -80 °C.

• Pseudomonas aeruginosa

  Material

  Table: TSA agar plate (for 1 liter)

  Material	Quantity
  tryptone	15 g
  soy peptone	5 g
  NaCl	5 g
  agar	15 g

  Table: TSB liquid media (for 1 liter)

  Material	Quantity
  tryptone	15 g
  soy peptone	5 g
  NaCl	5 g

  Table: M63 (for 1 liter)

  Material	Quantity
  20% Casein amino acids	225 mL
  5x M63 stock	200mL
  MgSO4 solution	1mL
  20% Glucose stock	10mL

  Procedure

  1.Recovery

  • Wipe the ampoule with absorbent cotton moistened with 75% ethanol
  • Heat the tip of the ampoule in a flame, then add a small drop of sterile water onto the heated tip to initiate cracking.
  • Break off the cracked tip using sterile forceps.
  • Using a sterile pipette, add 0.3–0.5 mL of liquid medium into the ampoule and gently shake to dissolve the lyophilized cells into suspension.
  • Transfer the entire cell suspension to a test tube.
  • Incubate at 37 °C with shaking at 200 rpm overnight.

  2.Maintenance

  • Spread the revived single colony onto a TSA agar plate, seal the edges with parafilm, and store at 4 °C.

  3.Cultivation

  • Pick a single colony and inoculate it into 5 mL of TSB medium. Incubate overnight at 37°C with 220 rpm shaking.

  4.Preservation

  • Add 0.6 mL of overnight culture and 0.4 mL of 50% glycerol into a cryovial.
  • Mix thoroughly by vortexing or repeated inversion. Store the bacterial glycerol stock at -80 °C.

DNA Cloning Methods

• Oligo Annealing

  Materials

  • Single-stranded Oligonucleotides Solution (complementary)
  • ddH₂O

  Procedure

  1. Dilute the single-stranded oligonucleotides to 10 μM using ddH₂O (at least 10 μL per dilution).

  2. Mix 10 μL of each complementary single-stranded oligonucleotide in a PCR tube.

  3. Place the mixture in the thermal cycler and set the program according to the table below.

     Table: Typical program for an oligo annealing reaction.

     Step	Temperature	Time
     Denaturation	95℃	3min
     Annealing	65℃	20s
     Hold	12℃	-

  4. Store at -20°C until further use (no more than 2 weeks).

• PCR

  Materials

  • PCR Master Mix (2X) (including 2X DNA Polymerase, 2X Buffer with Mg2²⁺, 2X dNTP)
  • Nuclease-free Water
  • Forward Primer & Reverse Primer
  • Template DNA

  Procedure

  1. Reaction Mixture Preparation

  1. Thaw all PCR components (primer, template DNA, etc.) on ice and mix each components gently.

  2. Dilute the primer to 10 μM using ddH₂O (at least 10 μL per dilution).

  3. Prepare the PCR reaction mixture in PCR-tubes on ice according to the order in the table below.

     Table: Typical composition of a PCR reaction.

     Component	Final Concentration	Volume
     Nuclease-free Water	-	(21-x) μL
     Template DNA	1~10 ng	x μL
     Primer Mix (10μM each)	0.8 μM each	4 μL
     PCR Master Mix (2X)	1X	25 μL
     Total	-	50 μL

  4. Mix gently by pipetting or brief vortex, then spin down.

  2. PCR Amplification

  1. Place the prepared PCR reaction mixture in the thermal cycler and set the cycling program according to the table below.

     Table: Typical program for a PCR reaction.

     Step	Temperature	Setting
     Initial Denaturation	92℃	3 min
     Denaturation	92℃	30 sec
     Annealing	55℃	30 sec
     Extension	68℃	15~60 sec/kb
     Go to Step 2	-	30~35 cycles
     Final Extension	68℃	10 min
     Hold	4℃	∞

     • Tips: It is recommended to use an extension time of 15 sec/kb for amplicons within 6 kb, and 60 sec/kb for amplicons left over 6 kb.

  2. Store at -20°C until further use.

• In-fusion Assembly

  Materials

  • 2 × CE Mix V3 (from ClonExpress Ultra One Step Cloning Kit V3)
  • ddH₂O
  • Linearized Vector
  • Insert Fragments

  Procedure

  1. Measure the DNA concentration of the linearized vector and the insert fragments separately using Nanodrop®.

  2. Prepare the reaction mixture in PCR-tubes on ice according to the table below.

     Table: Typical composition of an in-fusion assembly reaction.

     Component	Volume
     Linearized Vector	X μL
     Insert Fragments (n≤10)	Y1+Y2+…+Yn μL
     2 × CE Mix V3	5 μL
     ddH2O	to 10 μL

     • X/Y is the amount of vector/insert calculated by formula:
     • Single-fragment homologous recombination: The optimal mass of vector = [0.02 × number of bp] ng (0.03 pmol) The optimal mass of insert = [0.04 × number of bp] ng (0.06 pmol)
     • Multi-fragment homologous recombination: The optimal mass of vector = [0.02 × number of bp] ng (0.03 pmol) The optimal mass of each insert = [0.02 × number of bp] ng (0.03 pmol)
     • The recommended amounts of vector and insert are between 10~100 ng, respectively. When the optimal mass calculated using the formula is out of this range, the minimum/maximum input amount can be used directly.

  3. Mix gently by pipetting (do not vortex), then spin down.

  4. Place the reaction mixture in the thermal cycler and run the following program:

     • 50°C for 5-15 min (for single-fragment assembly) or 15-30 min (for multi-fragment assembly).
     • 4℃ forever.

  5. The recombination product can be used to transformation or stored at -20°C for one week.

• Agarose Gel Electrophoresis

  Materials

  • Agarose
  • 50× TAE
  • M5 GelRed Plus Nucleic Acid Dye (10,000×)
  • 6× loading buffer
  • DNA Marker (DL2000/DL2000 plus/DL15000…)
  • ddH₂O

  Procedure

  1. Gel Casting

  1. Prepare agarose gels according to the table below.

     Table: Typical formula for an agarose gel.

     Component	Final Concentration	Volume	Volume
     Agarose	0.8% (for plasmids or vectors) 1% (for insert fragments)	0.16 g 0.2 g	0.32 g 0.4 g
     50× TAE	1×	400 μL	800 μL
     ddH2O	-	19.6 mL	39.2 mL
     Total	-	20 mL	40 mL

  2. Microwave until fully dissolved (for ~2 min on the defrost setting).

  3. Once the solution has cooled down to about 60°C, add the M5 GelRed Plus Nucleic Acid Dye (10,000×) according to the dilution ratio and mix well.

  4. Pour the gel into the pre-assembled casting tray, then place a well comb and wait for ~30 min until the gel becomes firm.

  2. Electrophoresis

  1. For each sample, prepare the loading mix according to the table below.

     Table: Formula for the loading mix.

     Component	Volume	Volume
     DNA Sample	5 μL	25 μL
     6× Loading Buffer	1 μL	5 μL
     Total	6 μL	30 μL

     • Suggestions: For analytical electrophoresis (e.g., plasmid verification), load 6 μL. For preparative electrophoresis (e.g., PCR product purification), load 30 μL.

  2. Load DNA marker and the sample mixtures into the wells, respectively.

     • Choose a DNA marker whose range covers the sizes of the DNA fragments.

  3. Run gel at 150 V for 30 min, then observe the results in the gel imager.

• Gel Extraction

  Note: This protocol is effective for extracting DNA shorter than 10kb. The efficiency of extracting longer fragments is significantly decreased.

  Materials

  • Beyotime™ DNA Gel Extraction Kit:
    • Binding Buffer
    • Wash buffer (added ethanol)
    • Elution Buffer
    • DNA Purification Columns & Collection Tubes

  Procedure

  1. Gel Melting

  1. Excise the gel with intended DNA band and weigh the gel strip in a 1.5 mL EP tube. Break the strip if necessary.
  2. Add an equal volume of binding buffer to the gel fragments (e.g., add 100 μL of binding buffer for every 100 mg of gel). Mix thoroughly by vortex or inverting the tube.
  3. Incubate the mixture at 50-60°C for ~10 min until the gel is completely dissolved. Vortex or invert the tube 3-4 times during incubation to facilitate dissolution.
     • Note: After complete dissolution, continue incubating at 50-60°C for an additional 2 minutes. For DNA fragments >5 kb, use inversion instead of vortex.
  4. Transfer the dissolved gel solution to a DNA purification column and let it stand at room temperature for 1 min.
     • Note: If the volume exceeds the column capacity, load the sample in sequential steps: after centrifuging an initial portion, add the remaining solution to the same column and repeat.
  5. Centrifuge at ~16,000×g for 1 min. Discard the flow-through from the collection tube.

  2. Washing

  1. Add 700 μL of wash buffer to the column. Incubate at room temperature for 1 min.
  2. Centrifuge at ~16,000×g for 1 min to remove impurities. Discard the flow-through.
  3. Add 500 μL of wash buffer to the column. Centrifuge at ~16,000×g for 1 min to further remove impurities. Discard the flow-through.
  4. Centrifuge at ~16,000×g for an additional 1 min to eliminate residual ethanol and ensure complete evaporation.

  3. Elution

  1. Place the column in a clean 1.5 mL EP tube. Apply 30 μL of elution buffer directly to the center of the column. Then let it stand for 1 min.
  2. Centrifuge at ~16,000×g for 1 min. The eluate contains the intended DNA.

• Plasmid Extraction (by Magnetic Beads)

  Materials

  • BeyoMag™ Plasmid Mini Preparation Kit with Magnetic Beads
    • RNase A (add to Solution I before first use)
    • Solution I (Resuspension buffer)
    • Solution II (Lysis buffer)
    • Solution III (Binding/Neutralization buffer)
    • Solution IV (Wash buffer; add specified volume of anhydrous ethanol before first use)
    • Solution V (Elution buffer)
    • BeyoMag™ magnetic beads (mix well before use)
    • Magnetic separation rack (recommended: 12- or 24-well magnetic rack)

  Procedure

  1. Pellet cells

  • Transfer 1.5 mL of an overnight E. coli culture into a 1.5 mL microcentrifuge tube. Centrifuge at 5,000×g at room temperature for 1 minute to pellet the cells. Discard the supernatant.
  • Repeat once with another 1.5 mL sample so the tube contains pellet from a total of 3 mL overnight culture.
    Note: Typically LB overnight (~16 h) growing to an OD₆₀₀ of ~2–4 is recommended. If pellets are not compact, slightly increase centrifugation time. Do not centrifuge for too long or at too high speed — overly tight pellets are difficult to resuspend after adding Solution I.
  • After removing the supernatant, invert the open tube on absorbent paper to drain residual liquid. If cell density is low, consider using more culture (repeat pelleting 1–2 additional times). For high‑copy plasmids do not exceed ~3 mL culture per prep; for low‑copy plasmids do not exceed ~5 mL.

  2. Resuspend cells

  • Add 200 µL Solution I (Resuspension buffer containing RNase A) to each pellet. Resuspend thoroughly until no visible clumps remain. Vortex briefly or pipette up and down to break up the pellet.
    Note: Ensure RNase A is present in Solution I. The suspension should be homogeneous and turbid when held to a bright light.

  3. Alkaline lysis

  • Add 200 µL Solution II (Lysis buffer) to each tube. Immediately invert the tube gently 4–6 times to mix until the solution becomes clear.
    
    • Do NOT vortex. Vigorous mixing will shear genomic DNA and can contaminate the plasmid preparation. If clumps remain after 4–6 inversions, invert an additional 3–5 times and incubate at room temperature for 2–3 minutes. Total lysis time should not exceed 5 minutes.

  4. Neutralization

  • Add 200 µL Solution III (Binding/Neutralization buffer) to each tube and invert gently 4–6 times. A white flocculate (cellular debris) should appear.
    
    • Do NOT vortex. Excessive mixing can reduce plasmid quality.

  5. Clarify lysate by centrifugation

  • Centrifuge the tubes at maximum speed (≈13,000 rpm) at room temperature for 10 minutes. A white pellet containing precipitated debris will form.
  • While centrifuging, prepare a new labeled 1.5 mL tube for the cleared lysate.

  6. Bind plasmid to beads

  • Carefully transfer the cleared supernatant to a new 1.5 mL tube without disturbing the white pellet.
  • Add 50 µL well-mixed BeyoMag™ magnetic bead suspension to the supernatant (mix beads thoroughly before use). Gently mix by inversion or slow pipetting and incubate at room temperature for 3–5 minutes to allow plasmid binding.
    Note: If higher yield is required, increase bead volume or extend binding time.

  7. Separate beads on magnetic rack

  • Place the tube on a magnetic rack and wait until beads fully collect at the tube wall (typically <1 minute). Carefully remove and discard the supernatant without disturbing the bead pellet.

  8. Wash #1

  • Remove the tube from the rack. Add 750 µL Solution IV (wash buffer) and gently resuspend the beads by flicking or gentle inversion (do not vortex). Place the tube back on the magnetic rack and allow the beads to collect. Carefully remove and discard the supernatant.
    Note: If beads stick to the tube walls, briefly invert the tube while holding it on the rack so beads fall; then remove supernatant.

  9. Wash #2

  • Repeat the wash step with 450 µL Solution IV. Resuspend beads gently, magnetically separate, and remove as much residual liquid as possible.

  10. Dry beads

  • Air-dry the bead pellet to remove residual ethanol: leave the open tube at room temperature for 5–10 minutes or place in a 37°C dry oven for 5 minutes to evaporate traces of ethanol. Do not overdry.

  11. Elute plasmid

  • Add 50–100 µL Solution V (Elution buffer or nuclease-free water, pH ≥ 6.5) to the bead pellet. Gently resuspend the beads and incubate at room temperature for 3–5 minutes; gently invert or flick the tube 1–2 times during incubation to improve elution.
  • Place the tube on the magnetic rack until beads collect. Carefully transfer the supernatant (eluted plasmid) to a new labeled 1.5 mL tube. Store at -20°C.
    Note: To increase concentration use a smaller elution volume (e.g., 50 µL). Elution at 50–55°C can increase yield slightly compared with room temperature elution.

  Notes and troubleshooting

  • Always ensure RNase A is present in Solution I before starting.
  • Avoid vortex after adding Solution II and Solution III — mechanical shearing increases genomic DNA contamination.
  • If the pellet after neutralization is not compact or the lysate is cloudy, increase initial centrifugation time or combine more culture to increase biomass.
  • If you observe low yield or impurities, consider increasing bead volume, extending the binding time, and ensuring ethanol was added to Solution IV and that the final dry step was performed.
  • Bead aggregation: reduce culture input volume, ensure beads are fully resuspended before use, and thoroughly resuspend beads during washes.

• Plasmid Miniprep (by Spin‑column)

  Materials

  • Beyotime® Plasmid Mini Preparation Kit for All Purpose, Plasmid Miniprep Kit for All Purpose
    • RNase A (Solution I)
    • Solution I (Resuspension buffer)
    • Solution II (Lysis buffer)
    • Solution III (Neutralization buffer)
    • Solution PB (optional, column conditioning/wash buffer for EndA+ strains)
    • Solution IV (Wash buffer)
    • Solution V (Elution buffer)

  Procedure

  1. Pellet cells

  • Transfer 1.5 mL of an overnight E. coli culture into a 1.5 mL microcentrifuge tube. Centrifuge at 5,000 × g (approximately 5,000 rpm on many tabletop centrifuges) at room temperature for 1 minute to pellet the cells. Discard the supernatant.
  • Repeat once with another 1.5 mL sample so that each tube contains the pellet from a total of 3 mL overnight culture.
    Note: Typically LB overnight (~16 h) growing to an OD600 of ~2–4 is recommended. If pellets are not compact, slightly increase centrifugation time. Do not centrifuge for too long or at too high speed — overly tight pellets are difficult to resuspend after adding Solution I.
  • After removing the supernatant, add ~1.5 mL culture and repeat if needed. Invert the open tube on absorbent paper to drain residual liquid. If cell density is low, use more culture (repeat pelleting 1–2 additional times) to reach the desired biomass. For high‑copy plasmids do not exceed ~5 mL of culture per prep; for low‑copy plasmids do not exceed ~10 mL.

  2. Resuspended cells

  • Add 250 µL Solution I (resuspension buffer containing RNase A) to each pellet. Resuspend thoroughly until no visible clumps remain. Vortex at high speed for 5–10 seconds (or longer if required) to suspend the pellet.
  • If no vortex is available, pipette up and down or flick the tube to break up the pellet. The suspension should be homogeneous and turbid when held to a bright light.

  3. Alkaline lysis

  • Add 250 µL Solution II (lysis buffer) to each tube. Immediately invert the tube gently 4–6 times to mix until the solution becomes clear.
    • Do NOT vortex. Vigorous mixing will shear genomic DNA and can contaminate the plasmid preparation. If clumps remain after 4–6 inversions, invert an additional 3–5 times and incubate at room temperature for 2–3 minutes. Total lysis time should not exceed 5 minutes.

  4. Neutralization

  • Add 350 µL Solution III (neutralization buffer) to each tube and invert gently 4–6 times. A white flocculate (cellular debris) should appear.
    • Do NOT vortex. Excessive mixing can reduce plasmid quality.

  5. Clarify lysate by centrifugation

  • Centrifuge the tubes at maximum speed (~13,000 rpm) at room temperature for 10 minutes. A white pellet containing precipitated debris will form.
  • While centrifuging, prepare spin columns in labeled collection tubes.

  6. Bind plasmid to column

  • Carefully transfer the cleared supernatant to the spin column without disturbing the white pellet. Alternatively, aspirate the supernatant and apply it to the column.
  • Centrifuge at maximum speed for 30–60 seconds. Discard the flow‑through and retain the collection tube for reuse.

  7. Optional: PB wash (recommended for EndA⁺ strains)

  • Add 500 µL Solution PB to the column and centrifuge 30–60 seconds. Discard the flow‑through.
    • Note: This step is recommended (or required) when extracting plasmid from EndA+ strains (e.g., JM110, BL21(DE3), TG1, HB101) or strains with high nuclease activity or particular surface modifications. For common EndA‑ strains such as DH5α and XL1‑Blue, and if plasmid will be used for routine restriction digestion, PCR or cloning, this step can be skipped. If plasmid is intended for transfection, include this step to reduce nuclease contamination.

  8. Wash column

  • Add 750 µL Solution IV (wash buffer, typically contains ethanol) to the column. Centrifuge at maximum speed for 30–60 seconds. Discard the flow‑through and keep the collection tube.

  9. Dry column

  • Centrifuge the column (with the empty collection tube) at maximum speed for 1 minute to remove residual wash buffer and allow trace ethanol to evaporate. It is important to discard the flow‑through before this spin so residual wash is removed thoroughly.

  10. Elute plasmid

  • Place the spin column into a clean, labeled 1.5 mL microcentrifuge tube. Add 50 µL Solution V (elution buffer, or nuclease‑free water pH ≥ 6.5) directly to the center of the column membrane. Incubate for 1 minute at room temperature to allow buffer to soak into the membrane.
  • If higher concentration is desired, elute with a smaller volume (e.g., 35 µL) or perform a second elution and pool.

  11. Collect eluate

  • Centrifuge at maximum speed for 1 minute. The eluate contains the purified plasmid DNA.
    • Yield and concentration: typical plasmid concentration is ~0.1–0.3 mg/mL. To concentrate the DNA further, perform ethanol precipitation.

  Notes and troubleshooting

  • Always ensure RNase A is present in Solution I before starting.
  • Avoid vortexing after adding Solution II and Solution III — mechanical shearing increases genomic DNA contamination.
  • If the pellet after neutralization is not compact or the lysate is cloudy, increase initial centrifugation time or combine more culture to increase biomass.
  • Do not overload the column with excessive lysate; follow kit recommendations for maximum culture volume (high‑copy ≤ 5 mL, low‑copy ≤ 10 mL per column).
  • If you observe low yield or impurities, consider performing the optional Solution PB wash (Step 7) and ensure ethanol was present in Solution IV and that the final dry spin (Step 9) was performed.

  Table: Examples for EndA^- and EndA⁺ E. coli strains

  EndA-	EndA+
  BJ5183	BL21(DE3)
  DH1	CJ236
  DH20	HB101
  DH21	JM83
  DH5α	JM101
  JM103	JM110
  JM105	LE392
  JM106	MC1061
  JM107	NM522 (all NM series are EndA+)
  JM108	NM554
  JM109	P2392
  MM294	PR700 (all PR series are EndA+)
  SK1590	Q358
  SK1592	RR1
  SK2267	TB1
  SRB	TG1
  TOP10	Y1088 (all Y10 series are EndA+)
  XL1-Blue	BMH 71-18
  XLO	ES1301

• Genomic DNA Extraction

  Materials

  • RNase A
  • Proteinase K
  • Lyticase (yeast cell wall degrading enzyme)
  • Reducing agent (DTT or β-mercaptoethanol)
  • Sorbitol Buffer
  • Solution A
  • Solution B
  • Wash Buffer (ethanol must be added before use)
  • Elution Buffer (or nuclease-free water, pH ≥ 8.0 recommended)
  • Spin columns (DNA binding columns)
  • Collection tubes (1.5 mL or 2 mL microcentrifuge tubes)

  Procedure

  1. Pellet cells

  • Start with up to 5 × 10⁷ yeast cells. Centrifuge at 8,000 rpm for 1 min and remove the supernatant as completely as possible.

    Note: An overnight yeast culture typically reaches OD600 ≈ 2.0. Collect cells by centrifugation and wash once with PBS to remove residual medium.

  2. Yeast cell wall digestion

  • Resuspend the cell pellet in 470 µL Sorbitol Buffer and mix thoroughly.
  • Add 25 µL lyticase and 5 µL reducing agent (e.g., DTT or β-mercaptoethanol). Mix by inversion until homogeneous.
  • Incubate at 30°C for 1–2 h. During incubation, gently invert the tube a few times to mix.
  • Centrifuge at 12,000 rpm for 1 min and discard the supernatant. Keep the pellet.

  3. DNA separation

  • Add 300 µL Solution A to the pellet and resuspend thoroughly.
  • Add 2 µL RNase A, mix by inversion, and incubate at room temperature for 10 min.
  • Add 220 µL Solution B and then add 20 µL Proteinase K. Mix by inversion until homogeneous.
  • Incubate in a 65°C water bath for 15–30 min. During incubation, invert the tube a few times to mix.
  • Centrifuge at 8,000 rpm for 30 s and carefully transfer the supernatant to a new tube.
    Note: Work quickly: after cooling to room temperature the lysate may become cloudy.

  4. DNA purification

  • Allow the lysate to cool to room temperature (if still warm), then add 375 µL absolute ethanol and mix thoroughly.
  • Transfer the mixture to a spin column placed in a collection tube. Centrifuge at 8,000 rpm for 1 min. Discard the flow-through and place the column back into the collection tube.
  • Add 600 µL Wash Buffer (confirm ethanol was added) to the column. Centrifuge at 12,000 rpm for 1 min. Discard the flow-through and return the column to the collection tube. Then repeat this step.
  • Centrifuge the column at 12,000 rpm for 2 min to remove residual wash buffer. Then open the column and place it at room temperature or in a 50°C dry oven for a few minutes to evaporate any remaining ethanol.
    Note: Removing residual ethanol is critical — residual ethanol can inhibit downstream enzymatic reactions (restriction digest, PCR, etc.).
  • Transfer the spin column to a clean microcentrifuge tube. Pre-warm Elution Buffer to 65°C and add 50–200 µL directly to the center of the column membrane (recommended volume 50 µL).
  • Incubate at room temperature for 5 min, then centrifuge at 12,000 rpm for 1 min to collect the eluate.
  • (Optional) To increase yield, add the eluate back onto the column and centrifuge at 12,000 rpm for 2 min.

  5. Storage

  • Store purified genomic DNA at -20°C to prevent degradation.

  Notes and troubleshooting

  • Store lyticase at -20°C.
  • Before use, add absolute ethanol to the Wash Buffer according to the label on each bottle (each bottle requires 45 mL of absolute ethanol to be added separately).
  • All centrifugation steps are performed at room temperature using a benchtop centrifuge.
  • Avoid repeated freeze–thaw cycles of samples; repeated freezing and thawing fragments DNA and reduces yield.
  • If any buffers show precipitates, dissolve them by warming in a 65°C water bath before use; this does not affect extraction performance.
  • If the spin column becomes clogged during centrifugation, increase the centrifugation time to clear the blockage.
  • Use at least 50 µL for elution; smaller volumes may reduce recovery efficiency. Elution buffer pH affects recovery — if using water, adjust pH to ~8.0 (e.g., with a small amount of NaOH). pH < 7.0 will reduce elution efficiency.
  • DNA fragment size depends on sample storage time and mechanical shearing during handling. Evaluate fragment size by agarose gel electrophoresis.
  • Measure DNA concentration and purity by spectrophotometry. Double‑stranded DNA: OD260 of 1.0 ≈ 50 µg/mL. Expected OD260/OD280 ratio: 1.7–1.9. If eluted in water, OD ratio may appear lower due to pH and ionic differences; this does not necessarily indicate low purity.

• E. coli Transformaton (DH5α and BL21(DE3))

  Material

  • E. coli DH5α/BL21(DE3) competent cells
  • Plasmids (dissolved)
  • LB Broth
  • LB Agar Plate (without/with antibiotic)

  Procedure

  1. Place the competent cells on ice or in an ice-water for ~5 min until thawed.
     
     • Caution: Use the cells within 10 min after thawing, or it will significantly reduce transformation efficiency.
  2. Add the plasmid (with the amount of 1~2 ng) to 100 μL of competent cells. Mix gently by flicking the bottom of the tube or swirling gently. Do not mix by pipetting up and down.
  3. Immediately place the tube back on ice and incubate for 30 min.
  4. Transfer the tube directly to a 42°C water bath and incubate for 45 sec. Place the tube back on ice for 2 min to cool down. Do not shake or mix.
  5. Add 900 μL of 37°C pre-warmed, antibiotic-free LB broth to the tube. Mix by inverting the tube several times.
  6. Incubate the tube at 37°C in a shaking incubator (150 rpm) for 1 h to allow for recovery.
  7. Centrifuge the tube at 5,000 × g for 1 min at room temperature. Carefully aspirate and discard 900~950 μL of the supernatant, leaving 50~100 μL. Gently resuspend the bacterial pellet in the remaining supernatant by pipetting.
  8. Plate a small amount (e.g., ~10 μL) of the resuspended cells onto an LB agar plate without antibiotic as a positive control. Plate the remainder of the resuspended cells onto an LB agar plate containing the appropriate selective antibiotic.
  9. Invert the plates and incubate them overnight at 37°C.

• Yeast Chemically Competent Cells Preparation and Transfection

  Materials

  • Yeast strain stored in 15–20% glycerol at -80°C
  • YPD agar plates
  • YPD liquid medium
  • Beyotime® Competent Cell Preparation and Transformation Kit for Saccharomyces Cerevisiae
    • Buffer A
    • Buffer B
    • Buffer C
    • DMSO
    • Carrier DNA (salmon sperm DNA, boiled and chilled; 10 µl per transfection)
  • Plasmid DNA (0.1–1 µg per transfection)

  Procedure

  1. Strain activation

  • Remove yeast glycerol stock from -80°C and streak onto a YPD agar plate. Invert and incubate at 30°C for 2–4 days to revive and isolate single colonies.

  2. Inoculation

  • Using a sterile inoculating loop or pipette tip, pick a single fresh yeast colony and inoculate 10 mL of YPD liquid medium in a sterile 50 mL conical tube or culture flask.

  3. Overnight culture

  • Incubate the 10 mL starter culture at 28–30°C with shaking at ~250 rpm overnight (~12–16 hours).

  4. Subculture to mid‑log phase

  • Transfer an appropriate volume of the overnight culture into 50 mL YPD so that the starting OD600 is between 0.2–0.3 (typically transfer ~1–2 mL into 50 mL, adjust as needed). Incubate at 30°C with shaking at 250–300 rpm for 2–3 hours until OD600 reaches 0.4–0.6.

  • Note: The target OD₆₀₀ (0.4–0.6) is critical — cells in mid‑log phase give the best competence. As a rule of thumb, each 10 mL of culture can yield four 100 µL competent aliquots; scale accordingly.

  5. Preparation of competent cells

  • When the culture reaches OD600 = 0.4–0.6, transfer the appropriate volume (for example, 10 mL for a small prep) to a sterile 15 mL tube and centrifuge at 3,000 × g for 3 minutes at room temperature. Carefully discard the supernatant and keep the cell pellet.

  • Resuspend the pellet thoroughly in 7 mL Buffer A. Centrifuge again at 3,000×g for 3 minutes at room temperature and discard the supernatant.

  • Resuspend the washed pellet in 400 µL Buffer B. Incubate at room temperature for 10 minutes. After this incubation the cells are chemically competent and ready for transfection.

  6. Yeast transfection

  • To a sterile 1.5 mL tube containing 100 µL competent cells add 0.1–1 µg plasmid DNA and 10 µL heat‑denatured carrier DNA (ssDNA). Gently mix by pipetting or flicking the tube.

  • Add 600 µL Buffer C to the tube and mix by inverting or gently pipetting. Incubate in a 30°C water bath for 30 minutes.

  • After incubation, add 70 µL DMSO to the mixture and mix gently by inversion. Heat shock in a 42°C water bath for 15 minutes, then immediately transfer the tubes to an ice bath for 3–5 minutes.

  • Centrifuge the tubes at 12,000–14,000 × g for 10 seconds at room temperature. Carefully remove and discard the supernatant, then resuspend the cell pellet in 100 µL Buffer A.

  • Plate the entire resuspended volume onto the appropriate selective agar plate. Invert and incubate at 30°C for 2–3 days until single colonies appear.

  Notes and Troubleshooting

  • Immediately aliquot competent cells into 100 µL portions in sterile 1.5 mL tubes for transfection.
  • The procedure above is written for a typical small-scale preparation where 10 mL of culture yields four 100 µL aliquots of competent cells. Scale volumes proportionally for larger or smaller preparations.
  • Prepared competent cells are best used immediately. Short-term storage at 4°C for up to 1 week is possible but efficiency will decrease with time. For longer storage, add glycerol to a final concentration of 15% and freeze at -80°C; frozen aliquots are effective for at least ~3 months but efficiency may decline.
  • Factors affecting transfection efficiency:
    • Yeast strain differences and growth conditions strongly influence efficiency.
    • Using highly pure plasmid DNA improves efficiency; different plasmid backbones and sizes will transform at different rates.
    • Adding DMSO prior to the 42°C heat step can increase transfection efficiency by 2–5× in many cases.
  • If transfection efficiency is low:
    • Verify cells were in mid‑log phase (OD600 0.4–0.6) at the time of preparation.
    • Confirm carrier DNA was properly denatured (boiled, then chilled on ice) and is present in the transfection mix.
    • Ensure reagents — especially LiAc or PEG if present in your Buffers — are fresh and prepared correctly.

• Yeast Electrocompetent Cells Preparation and Transfection

  Materials

  • 1 M DTT (Dithiothreitol)
  • 1 M HEPES buffer (pH 8.0)
  • 1 M Sorbitol
  • YPD plates and YPD liquid medium
  • 1.5 mL and 50 mL sterile centrifuge tubes
  • Sterile microcentrifuge tubes (1.5 mL)
  • Sterile pipettes and tips
  • 0.2 cm electroporation cuvettes (ice pre-chilled)
  • Electroporator capable of settings below
  • Ice bucket and pre-chilled solutions

  Procedure

  1. Growth / Activation

  • Pick a single freshly isolated colony from a YPD plate and inoculate 5–10 mL YPD liquid medium.
  • Incubate overnight at 30°C, 220 rpm.
  • Next day dilute the overnight culture 1:100 into 100 mL fresh YPD.
  • Grow at 30°C, 220 rpm for ~3–5 hours until OD600 reaches 0.8–1.0 (log phase).

  2. Pretreatment

  • Transfer the culture into sterile centrifuge tubes and pellet cells at 4°C, 5,000 × g for 1 min.
  • Remove the supernatant. Gently resuspend the pellet in 25 mL ice-cold sterile water.
  • Centrifuge again at 4°C, 5,000 × g for 1 min and discard the supernatant.
  • Add 10 mL freshly prepared pretreatment buffer (prepare immediately before use):
    • 9.8 mL sterile water
    • 100 µL 1 M HEPES (pH 8.0)
    • 100 µL 1 M DTT
  • Gently resuspend the cells, transfer to a 50 mL tube, and incubate with gentle shaking (≈100 rpm) at 30°C for 15 minutes.

  3. Wash and concentrate

  • After incubation, pellet cells at 4°C, 5,000 × g for 1 min. Discard the supernatant.
  • Resuspend the pellet gently in 25 mL ice-cold 1 M sorbitol. Centrifuge at 4°C, 5,000 × g for 1 min.
  • Repeat the sorbitol wash once more. After the second wash, carefully remove all residual liquid by pipetting.
  • Resuspend the cells in 0.5 mL ice-cold 1 M sorbitol.
  • Use the prepared competent cells immediately (do not store).

  4. Electroporation setup

  • Switch on the electroporator and set the following parameters:
    • Voltage: 1.5 kV
    • Capacitance: 25 µF
    • Resistance: 200 Ω
  • Place a chilled 0.2 cm electroporation cuvette and a 1 mL tube containing ice-cold 1 M sorbitol on ice.

  5. Mix DNA and cells

  • Use an ice-cold 1.5 mL microcentrifuge tube.
  • Add 40–50 µL freshly prepared competent cells to the tube.
  • Add 200–500 ng plasmid DNA (keep volume minimal, < 5 µL if possible). Mix by gentle pipetting. DO NOT VORTEX.
  • Immediately transfer the mixture to the bottom of the pre-chilled electroporation cuvette, ensuring the liquid covers the metal surface and no bubbles are present.

  6. Pulse (electroporation)

  • Wipe any moisture from the outside of the cuvette with a tissue and place it in the electroporator sample holder.
  • Deliver the pulse by pressing the instrument’s pulse button. Be aware of high-voltage risk.
    • A successful pulse is usually accompanied by an audible beep and a time constant (τ) readout. Optimal τ is typically between 4.5–5.5 ms.
  • Immediately add 1 mL ice-cold 1 M sorbitol to the cuvette. Gently pipette up and down a few times to resuspend the cells and transfer the mixture to a new 1.5 mL tube. Work quickly to recover damaged cells.

  7. Recovery

  • Add an equal volume (1 mL) of 2× YPD medium to the cell suspension to give a final rich recovery medium (YPDS).
  • Incubate at 30°C with shaking at 150 rpm for 1–2 hours to allow expression of selectable markers and recovery.

  8. Plating

  • Pellet cells gently (3,000 × g, 3 min). Remove most of the supernatant and leave ~100–200 µL to resuspend the cells.
  • Plate 100 µL of the cell suspension evenly onto the appropriate selective SD plates. You may plate serial dilutions (e.g., undiluted, 1:5, 1:10) to estimate transformation efficiency.

  9. Incubation and colony appearance

  • Invert plates and incubate at 30°C for 2–4 days until colonies are visible. Larger plasmids often give rise to slower-growing transformants.

  Notes and troubleshooting

  • Keep everything cold: Except for the DTT treatment (pretreatment at 30°C) and the recovery step, perform all centrifugation, washing and handling steps on ice or at 4°C.
  • Cell state: Ensure yeast are in mid-log phase (OD600 = 0.8–1.0) for optimal competency.
  • Time constant (τ): Monitor the electroporation time constant after pulsing:
    • τ < 4 ms: possible air bubbles in the sample or insufficient voltage.
    • τ > 6 ms or a loud arcing / crackling sound: indicates high conductivity (residual salts) in the sample. Increase wash stringency to reduce salts.
  • Plasmid size: Transformation efficiency decreases with increasing plasmid size. For plasmids >30 kb consider increasing DNA amount (up to ~1 µg) and further optimization.
  • Work quickly after pulse: Adding sorbitol immediately after pulsing is critical to rescue electro-damaged cells.
  • Sterility: Use sterile reagents, tips and tubes to minimize contamination during recovery and plating.

Protein Expression & Analysis

• Bacterial Protein Induced Expression & Extraction

  Materials

  • Ampicillin stock solution
    • 100 mg/ml aqueous solution
  • 1M IPTG stock solution
  • Buffer W
    • 100 mM Tris/HCl (pH 8)
    • 150 mM NaCl
    • 1 mM EDTA
  • 5× SDS-PAGE sample buffer
    • 0.25 M Tris·Cl (pH 8.0)
    • 25% glycerol
    • 7.5% SDS
    • 0.25 mg/ml bromophenol blue
    • 12.5% v/v β-mercaptoethanol

  Table: LB liquid media (for 1 liter)

  Material	Quantity
  Yeast Extract	5 g
  Tryptone	10 g
  NaCl	10 g

  Procedure

  • Inoculate 10 mL of LB medium containing 100 μg/mL ampicillin, and incubate overnight at 37°C with shaking at 200 rpm.

  • Dilute the overnight culture 1:100 into 100 mL of LB medium supplemented with ampicillin, and continue growth at 37°C with shaking at 200 rpm.

  • Induce protein expression when the OD₆₀₀ reaches 0.5–0.6.

  • Add 50 μL of IPTG to achieve a final concentration of 0.5 mM, and induce protein expression for 18 hours at 25°C with shaking at 180 rpm.

• Yeast Protein Expression & Extraction

  Yeast Protein Induced Expression

  Materials

  • 100× Histidine Solution
    • 0.2g Histidine
    • 100mL ddH₂O
  • 100× Methionine Solution
    • 0.2g Methionine
    • 100mL ddH₂O
  • 100× Leucine Solution
    • 1g Leucine
    • 100mL ddH₂O
  • 100× Uracil Solution
    • 0.2g Uracil
    • 100mL ddH₂O
  • 30% galactose-raffinose mixture (Gal:Raf=2:1)

  Table: Stock solution (all sterilized by filtration)

  Stock	Composition (for 1 liter)
  40% Galactose Stock	400.0 g galactose
  20% Raffinose Stock	200.0 g raffinose
  20% Glucose Stock	200.0 g glucose
  Amino Acid & Nucleotide Stock	100 mg L-Leucine/20 mg L-Histidine/20 mg L-Methionine/20 mg Uracil

  Table: SC-Ura-Leu-Met-His media (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium Sulfate	5 g
  DO Supplement	0.6 g

  Table: SC-Ura media (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium Sulfate	5 g
  All 3 Amino Acid Stock	10 mL each
  DO Supplement	0.6 g

  Table: SC-Leu media (for 1 liter)

  Material	Quantity
  YNB	1.7 g
  Ammonium Sulfate	5 g
  Ura, Met, His Stock	10 mL each
  DO Supplement	0.6 g

  Table: SD-Ura activation media

  Material	Quantity
  SC-Ura Liquid Media	45 mL
  20% Glucose Stock	5 mL

  Table: SD-Leu activation media

  Material	Quantity
  SC-Leu Liquid Media	45 mL
  20% Glucose Stock	5 mL

  Table: SC-Ura+Raf scale-up media

  Material	Quantity
  SC-Ura Liquid Media	450 mL
  20% Raffinose Stock	50 mL

  Table: SC-Leu+Raf scale-up media

  Material	Quantity
  SC-Leu Liquid Media	450 mL
  20% Raffinose Stock	50 mL

  Note: All media preparations requiring sterilization are autoclaved; filter sterilization is performed under biosafety cabinet for heat-sensitive components.

  Procedure

  1. Activation & Seed Culture

  • Pick a single positive colony (verified by electrophoresis) and inoculate it into 10 mL of the corresponding activation medium.

  2. Expansion & Induction

  • Conventional induction method
    • Measure the OD₆₀₀ of the overnight culture.
    • Inoculate an appropriate volume of the culture into 100 mL of the corresponding scale-up medium to achieve an initial OD₆₀₀ between 0.2 and 0.4.
    • Incubate at 30°C with shaking at 250 rpm, and monitor OD₆₀₀ every hour.
    • When OD₆₀₀ reaches between 0.8 and 1.2, add 5 mL of a 40% galactose solution to induce protein expression.
    • Induce at 30°C with shaking at 250 rpm for the desired duration.
  • One-step induction method
    • Measure the OD₆₀₀ of the overnight culture.
    • Inoculate an appropriate volume of the culture into 100 mL of the corresponding scale-up medium to achieve an initial OD₆₀₀ between 0.2 and 0.4.
    • Add 10 mL of the 30% galactose-raffinose mixture.
    • Induce at 30°C with shaking at 250 rpm for the desired duration.

  Yeast Protein Constitutive Expression

  Materials

  • 100× Histidine Solution
    • 0.2g Histidine
    • 100mL ddH₂O
  • 100× Methionine Solution
    • 0.2g Methionine
    • 100mL ddH₂O
  • 100× Leucine Solution
    • 1g Leucine
    • 100mL ddH₂O
  • 100× Uracil Solution
    • 0.2g Uracil
    • 100mL ddH₂O

  Table: Stock solution (all sterilized by filtration)

  Stock	Composition (for 1 liter)
  20% Glucose Stock	200.0 g Glucose

  Table: YPD (for 500 mL)

  Material	Quantity
  BeyoPure™ YPD Broth premixed powder (namely, 5 g/500mL yeast extract, 10 g/500mL peptone, 10 g/500mL D-glucose)	1 bottle
  agar for plates	10 g

  Table: SC-His media

  Material	Quantity
  YNB	1.7 g
  Ammonium Sulfate	5 g
  Ura, Leu, Met Stock	10 mL each
  DO Supplement	0.6 g

  Table: SC-His-Ura media

  Material	Quantity
  YNB	1.7 g
  Ammonium Sulfate	5 g
  Leu, Met Stock	10 mL each
  DO Supplement	0.6 g

  Table: SC-His-Ura-Leu media

  Material	Quantity
  YNB	1.7 g
  Ammonium Sulfate	5 g
  Met Stock	10 mL each
  DO Supplement	0.6 g

  Procedure

  1. Activation & Seed Culture (Using Drop-out Medium)

  • Pick a single positive colony (verified by electrophoresis) or inoculate 10 µL of liquid culture into 10 mL of the corresponding SC Selection + Glc activation medium.
  • Incubate at 30°C, 250 rpm overnight (16-18 hours).

  2. Expansion Culture & Expression (Using YPD Medium)

  • Measure the OD₆₀₀ of the overnight culture.
  • Dilute the culture 1:20 to 1:50 into 200 mL of corrensbonding expansion medium to achieve an initial OD₆₀₀ between 0.2 and 0.4.
  • Induce at 30°C with shaking at 250 rpm for the desired duration.

  Yeast Protein Extraction

  Intracellular Protein Extraction

  Materials

  • Yeast Protein Extraction Kit
    • Isotonic solution
    • Hypotonic solution
    • Chymotrypsin Inhibitor
    • PMSF
  • β-Mercaptoethanol

  YPD medium (Per liter)

  Component	Amount
  Yeast Extract	10g
  Peptone	20g
  Dextrose(D-glucose)	20g

  Procedure

  • Transfer single colonies from the selective medium to 100 mL of YPD medium and culture at 30°C until the logarithmic growth phase ( OD₆₀₀ = 0.8–1.2, approximately 15 hours). Centrifuge at 8000 rpm (5000 × g) for 1 minute, collect the cells, and measure the wet weight.

  • For every 50 mg of wet cell weight, add 500 μL of isotonic solution, 5 μL of snailase, and 1 μL of β-mercaptoethanol. Resuspend the cells by pipetting or vortexing.

  • Incubate in a 30°C water bath for 60 minutes, then centrifuge at 5000 rpm (2500 × g) for 1 minute. Discard the supernatant and collect the pellet.

  • Resuspend the pellet in 500 μL of isotonic solution, then centrifuge at 5000 rpm (2500 × g) for 1 minute. Discard the supernatant and collect the pellet. Repeat this washing step once.

  • Resuspend the pellet in 500 μL of hypotonic solution (before use, add 5 μL of PMSF to 500 μL of hypotonic solution). Freeze the suspension in a -20°C freezer for 30 minutes, then thaw at room temperature (repeat this cycle 2–3 times).

  • Centrifuge the solution at 12,000 rpm (12,830 × g) for 5 minutes. The resulting supernatant is the yeast protein extract (store at -20°C).

  Cell Wall Protein Extraction - Mechanical Homogenization

  Materials

  • Western & IP Lysis Buffer
  • Grinding Beads

  Procedure

  • Centrifuge at 5000 × g for 3 minutes, remove the supernatant, and resuspend the pellet with 1 mL of pre-chilled PBS. Wash by repeating this step three times.

  • Resuspend the pellet in 1 mL of Western & IP cell lysis buffer, transfer to a sample tube, and add an equal volume of 0.1 mm glass beads.

  • Homogenize using a homogenizer at 6 m/s for 10 seconds per homogenization step, with a 10-second pause between steps, repeating for a total of 9 times.

  • After homogenization, take a small aliquot of the suspension and observe under a microscope to assess the extent of cell disruption. If insufficient, repeat the homogenization.

  • After homogenization, briefly centrifuge the suspension and transfer the supernatant to a 1.5 mL microcentrifuge tube.

  • Wash the grinding beads and sample tube, drain excess liquid, and return to their original positions.

  • Centrifuge at 16,000 × g for 10 minutes at 4°C.

  Cell Wall Protein Extraction - Enzymatic Lysis

  Materials

  • Yeast Protein Extraction Kit
    • Isotonic solution
    • Hypotonic solution
    • Chymotrypsin Inhibitor
    • PMSF
  • β-Mercaptoethanol
  • Lyticase

  Procedure

  • Collect cells by centrifugation (5,000 × g, 5 minutes). Discard the supernatant.
  • Measure the wet weight of the pellet.
  • For every 50mg of wet cell weight, add 500 μl of isotonic solution, 1.4 μl of β-mercaptoethanol, 7 μL of PMSF, and and 10,000 U of Lyticase (cell wall lysis enzyme, 200 U/mg). Resuspend the pellet by pipetting or vortexing.
  • Incubate at 37°C in a metal bath for 90 minutes. Gently invert the tube to mix every 30 minutes.
  • Centrifuge at 5,000 × g for 1 minute. Carefully collect the supernatant and transfer it to a new tube. Discard the pellet.
  • Centrifuge the supernatant at 12,000 × g for 10 minutes, carefully aspirate and transfer the supernatant, discarding the pellet. Label the supernatant and store at -20°C.

  Isolation of Secretory Protein

  Procedure

  • Aliquot the cell suspension into 50 mL centrifuge tubes, balance the tubes, and centrifuge at 5000 × g for 2 minutes to pellet the cells.
  • Carefully aspirate the supernatant and transfer it to a new centrifuge tube.
  • Repeat the above steps once.
  • Label the supernatant and store at -20°C.

• Purification

  Purification of His-tagged Proteins

  Materials

  • BeyoMag™ His-tagged Protein Purification Agarose Magnetic Beads (IDA-Ni)
  • Protein sample
  • 20% ethanol
  • Binding/Wash Buffer
  • Elution Buffer
  • 1X TBS
  • 2ml centrifuge tube
  • 0.22 μm pore size filter membrane
  • Syringe

  Binding/Wash Buffer

  Component	Concentration
  Tris	10mM
  NaCl	500mM

  Caution: Filter the ddH₂O and buffer solution through a 0.22 μm pore size filter membrane before use, and store at 4℃ for future use.

  Elution Buffer

  Component	Concentration
  Tris	10mM
  NaCl	500mM
  Imic	500mM
  Caution: Filter the ddH2O and buffer solution through a 0.22 μm pore size filter membrane before use, and store at 4℃ for future use.

  Procedure

  1. Preparation of Anti-His Magnetic Beads

  1. Resuspend the Magnetic Beads in the vial (gently pipette for 10 times, do not vortex). Transfer 10-20 μl of Magnetic Beads suspension into a new centrifuge tube (for 500 μl of protein sample). The amount of bead suspension can be scaled up or down proportionally based on the volume of protein sample.
     Caution: If using more than 0.2 ml of bead suspension, place the tube into a magnetic stand to collect beads and remove the supernatant.
  2. Add 500μl of 1X TBS (ST661/ST665) to the beads and pipette gently to mix. Place the tube into a magnetic stand to collect the beads against the side of the tube. Remove and discard the supernatant. Repeat this step twice.
  3. Resuspend the Magnetic Beads with 1X TBS at an equal volume to the initial volume of bead suspension taken in step 1.

  2. Protein Purification

  1. Add the protein sample to the washed magnetic beads and incubate them on a rotary mixer at room temperature for 20-30 minutes.
  2. Place the centrifuge tube on the magnetic stand for 10 seconds to separate. Gently aspirate and retain a portion of the supernatant for measurement of purification efficiency, and discard the remaining supernatant.
  3. Add 1.5 ml of Binding/Wash Buffer, gently vortex to resuspend the magnetic beads, perform magnetic separation for 10 seconds, then discard the supernatant.
  4. Repeat step 3 twice.
  5. Elution #1: Add 0.75 ml of Elution Buffer, gently invert the centrifuge tube to ensure the magnetic beads are suspended. Incubate at room temperature for 5 minutes, then perform magnetic separation for 10 seconds. Collect the eluate into a new centrifuge tube.
  6. Elution #2: Add another 0.75 ml of Elution Buffer to the magnetic beads and repeat the above steps. Combine the eluate from the two rounds.
  7. Mix the elution solution thoroughly and evenly distribute it into two 1.5 ml centrifuge tubes. Add an equal volume of 50% glycerol to each centrifuge tube and mix well.
  8. Divide the mixed liquid into 100 µl portions for each centrifuge tube and store at -20°C.

  3. Cleaning and Regeneration of Magnetic Beads

  1. Add 2 ml of Elution Buffer to the centrifuge tube, invert several times, vortex for 10 seconds to resuspend the magnetic beads, perform magnetic separation, and remove the supernatant. Repeat this process 3 times.
  2. Add 2 ml of 20% ethanol to the centrifuge tube, invert several times, vortex for 10 seconds to resuspend the magnetic beads, perform magnetic separation, and remove the supernatant. Repeat this process 3 times.
  3. Store the magnetic beads in 20% ethanol, ensuring the total volume is equal to the initial suspension volume. Place it at 4°C and it can be used for the purification of the same protein in the next experiment.

  4. Calculation of the purification efficiency and purity

  1. Calculate the purification efficiency using the following formula.
     $$\begin{align*} \text{Purification efficiency} &= \frac{\text{Total amount of purified protein}}{\text{Total amount of protein in sample}} \\ \text{Total amount} &= \text{Concentration (mg/ml)} \times \text{Volume (ml)} \end{align*}$$
  2. Separate the purified protein by SDS-PAGE. After electrophoresis, measure the gray-scale integrals of the target band and the total protein band by ImageJ. Calculate the proportion of the target protein in the total protein to obtain the purity.

  Purification of FLAG-tagged Proteins

  Materials

  • BeyoMag™ Anti-Flag Magnetic Beads
  • Protein sample
  • 1X TBS
  • 3X Flag Stock Solution (5mg/ml)

  Procedure

  1. Preparation of Anti-Flag Magnetic Beads

  1. Resuspend the Magnetic Beads in the vial (gently pipette for 10 times, do not vortex). Transfer 10-20 μl of Magnetic Beads suspension into a new tube (for 500 μl of protein sample). The amount of beads suspension can be scaled up or down proportionally based on the volume of the protein sample.
     Caution: If using more than 0.2 ml of bead suspension, place the tube into a magnetic stand to collect beads and remove the supernatant.
  2. Add 500μl of 1X TBS to the beads and pipette gently to mix. Place the tube into a magnetic stand to collect the beads against the side of the tube. Remove and discard the supernatant. Repeat this step twice.
  3. Resuspend the Magnetic Beads with 1X TBS at an equal volume to the initial volume of beads suspension taken in step 1.

  2. Protein binding

  1. Add 500 μl of cell lysate to the washed beads, pipette gently to resuspend beads, and incubate for 2 hours at room temperature or overnight at 4ºC while gently rotating the tube on a rotary mixer.
     Caution: Occasional aggregation of magnetic beads during the binding process doesn’t affect experimental results.
  2. Place the tube into a magnetic stand to collect beads against the side of the tube. Remove and discard the supernatant.
     Caution: A small amount of supernatant can be reserved in a clean EP tube for examination of the binding results.
  3. Wash beads with 500μl of 1X TBS and gently pipette to mix. Place the tube into a magnetic stand to collect the beads against the side of the tube. Remove and discard the supernatant. Repeat this step 3 times.
     Caution: The A₂₈₀ of supernatant can also be measured to determine whether the beads are washed thoroughly. Repeat washing until the A₂₈₀ is smaller than 0.05.

  3. Elution

  1. Dilute the 3X Flag Stock Solution at 5mg/ml with 1X TBS buffer to a final concentration of 150 μg/ml to obtain 3X Flag elution buffer.
  2. Add 100μl of 3X Flag elution buffer to resuspend the beads by pipetting gently and incubate with gentle shaking or on a rotator for 30-60 minutes at room temperature or 1-2 hours at 4ºC. The elution efficiency can be improved by prolonging the incubation time appropriately or by repeated elution.
  3. Place the tube into a magnetic stand for 10 seconds and transfer the supernatant to a clean EP tube.
  4. For immediate use, keep the eluates at 4ºC; or store at -20ºC or -80ºC for long-term storage.

  Purification of Myc-tagged Proteins

  Materials

  • BeyoMag™ Anti-Myc Magnetic Beads
  • Protein sample
  • 1X TBS
  • 2ml centrifuge tube
  • c-Myc polypeptide solution (150 μg/mL)

  Procedure

  1. Preparation of Anti-Myc Magnetic Beads

  1. Use a pipette to gently stir and resuspend the Anti-Myc magnetic beads. Based on the calculation that 20 μl of magnetic bead suspension is needed for every 500 μl of sample, take an appropriate amount of Anti-Myc magnetic beads and transfer them to a clean 2 ml centrifuge tube. Add 0.5 mL of 1X TBS.
  2. Use a pipette to gently stir and thoroughly resuspend the Anti-Myc magnetic beads. Place them on a magnetic stand and let them separate for 10 seconds. Remove the supernatant.
  3. Repeat step 2 twice.
  4. Resuspend the Anti-Myc magnetic beads with 1X TBS in an amount equivalent to the initial volume.

  2. Purification of the protein sample

  1. Add the corresponding volume of protein sample to the magnetic beads and place them on a rotary mixer. Incubate at room temperature for 2 hours.
  2. Place the sample on the magnetic stand and separate for 10 seconds. Gently aspirate and retain a portion of the supernatant for use in the subsequent purification effect detection, and discard the remaining supernatant.
  3. Add 500 μl of 1X TBS and gently vortex to resuspend the Anti-Myc magnetic beads with a pipette. Place the sample on a magnetic stand and separate for 10 seconds. Then remove the supernatant.
  4. Repeat step 3 twice.
  5. For every 20 μl of the original magnetic beads, add 100 μl of the c-Myc peptide elution solution (150 μg/ml), mix well, and then place it on a rotating shaker for incubation at room temperature for 30 to 60 minutes.
  6. Place the sample on a magnetic stand for 10 seconds for separation. Then transfer the supernatant to a new centrifuge tube.
  7. The supernatant is the purified Myc-tagged protein and should be stored at 4 degrees for future use, or at -20 degrees or -80 degrees for long-term storage.

• SDS-PAGE

  Materials

  • Ultra Pure Water (ddH₂O)
  • SDS-PAGE Gel One-step Super Quick Preparation Kit (Red Stacking Gel) (Beyotime, China)
    • 30% Acr–Bis (29:1)
    • 1M Tris–HCl, pH 8.8
    • 10% SDS (Sodium Dodecyl Sulfate)
    • Gel Polymerization Catalyst
    • 1M Tris–HCl, pH 6.8 (Red color)
  • Protein Sample
  • eStain^® L1 Protein Staining System

  Procedure

  1. Gel Casting

  • Use Beyotime SDS-PAGE Gel One-step Super Quick Preparation Kit (Red Stacking Gel) for gel casting.
  • Choose an appropriate acrylamide % for the gels depending on the molecular weight of the target protein according to.

  Table 1. The Optimal Separation range of SDS-PAGE gels at various concentrations

  Gel Percentage	MW of target protein (kDa)
  6%	50-150
  8%	30-90
  10%	20-80
  12%	12-60
  15%	10-40

  • Prepare a 10% Gel Polymerization Catalyst solution with ddH₂O or other high-purity water. For example, weigh 0.1 g of Gel Polymerization Catalyst, dissolve it in ultrapure water and make up to 1 mL.
    Caution: The gel polymerization catalyst solution is prone to degradation. It is recommended to aliquot and store at -20ºC, where it typically remains effective for six months.
  • Prepare 5 mL resolving gel mix and 2 mL stacking gel mix.
    Caution: Work carefully in order to avoid direct exposure to toxic acrylamide.

  Table: Resolving Gel

  Reagents	Volume (mL)
  SDS-PAGE One-step Resolving Gel Master Mix	5
  10% gel polymerization catalyst	0.05
  TEMED Substitute	0.002

  Note: The SDS-PAGE One-step Resolving Gel Master Mix should be matched to the MW of target protein in Table 1.

  Table: Stacking Gel

  Reagents	Volume (mL)
  SDS-PAGE One-step Resolving Gel Master Mix	5
  10% gel polymerization catalyst	0.05
  TEMED Substitute	0.002

  • Add 4.5 mL of the resolving gel mix, and then immediately add the stacking gel mix to fill the assembled glass plates. While adding, pay attention to avoiding bubbles, and the liquid surface should remain flat.
    Caution 1: Shake the Stacking Gel Master Mix to mix it well before use.
    Caution 2: When adding the stacking gel, it is important to gently and slowly add it with a 1mL pipette while moving it left and right, instead of pouring it in one spot. This is to prevent the stacking gel from rushing into the resolving gel and disrupting the smoothness of the gel interface.
  • Insert the comb and leave at room temperature (~25℃) until gels are fully solidified (usually within 10-30 minutes).

  2. Sample Preparation

  • According to experimental requirements, lyse cell or tissue samples with appropriate lysis buffers.
  • Protein quantification (optional). To ensure consistent protein loading of each sample, protein concentration of each sample needs to be determined. The compatibility of different protein quantification methods varies greatly for some detergents, reducing agents, or other factors.
  • Preheat the Thermal Block Heater to 95℃.
  • Add SDS sample buffer to the samples to yield a final concentration of 1X.
  • Vortex and briefly spin down samples.
  • Incubate the samples in the heat block for 5 mins at 95°C to fully denature proteins.
    Caution: Samples should always be heated at 95℃. Excessive temperatures (e.g., 100℃) or time (e.g., more than 15 minutes) may result in protein degradation or abnormal coloration of the indicator in the sample loading buffer.
  • After cooling to room temperature, the mixture can be loaded onto gels for electrophoresis.
    Caution: When using a water bath to dissolve BeyoWB™ SDS-PAGE sample loading buffer (5X), store it at room temperature immediately after dissolution to avoid prolonged exposure to the water bath. After use, it should be stored as soon as possible at -20ºC. If necessary, aliquot the loading buffer.

  3. Electrophoresis

  • Measure out 50 mL of BeyoWB™ SDS-PAGE Electrophoresis Buffer (Tris-Gly, 20X) provided in the kit and transfer it into a clean graduated cylinder. Add ultrapure water, deionized water, or distilled water to a final volume of 1L and mix thoroughly to obtain 1.0 L of BeyoWB™ SDS-PAGE Electrophoresis Buffer (Tris-Gly, 1X). The buffer can also be prepared in any required volume. Unused electrophoresis buffer can be stored at room temperature for 2-3 days or at 4℃ for 1-2 weeks.
  • Insert the gel into the running chamber. Secure the gel in the electrophoresis tank and slowly and gently remove the comb.
  • Fill the inner chamber with the electrophoresis buffer and ensure there are no leaks. In the outer chamber, add enough buffer to cover the bottom of the gel. Typically, 400 mL of electrophoresis buffer is sufficient for simultaneous running of two gels, and 800 mL for simultaneous running of four gels.
  • Load an appropriate volume of protein sample prepared. First, add marker into the first lane. And then add protein samples. Gently insert the pipette tip vertically into the sample well for loading. The optimal sample volume can be determined experimentally. Overloading the sample can lead to band distortion and overlay of strong signals. Typically, load 5-20 μL of samples, although the maximum sample volume for each well in the provided precast gel is 30 μL.
  • Close the lid of the electrophoresis tank and insert the power supply plugs into the corresponding power sockets of the electrophoresis unit.
    • If using precast gel, it is recommended to run the electrophoresis at 180 V for 25 minutes. However, the electrophoresis time may vary depending on the electrophoresis tank and ambient temperature. Generally, stop the electrophoresis when the bromophenol blue reaches near the bottom of the gel or when the target protein is expected to be properly separated according to the electrophoresis result of the prestained protein ladder.
    • If using the gel made in step 1, run the electrophoresis at 10 V until all the bands enter the separating gel. Then, run the electrophoresis at 150 V until the bromophenol blue reaches near the bottom of the gel.

  4.Gel Staining

  • Place the gel horizontally on the mesh surface of the gel fixation clamp. Spread a filter paper evenly on the gel, then close the gel fixation clamp.
  • Insert the gel fixation clamp into one channel of the staining machine and start the staining process.
  • When the process finishes, pull out the clamp and take the gel out for photographing.

• Western Blot

  Materials

  • BeyoWB™ 80 min Electrophoresis, Transfer and Western Blot Kit (Beyotime, China)
    • BeyoWB™ PVDF Membrane (6.6x8.5 cm, 0.45μm)
    • BeyoWB™ Transfer Buffer (10X)
    • BeyoWB™ BeyaECL A Solution
    • BeyoWB™ BeyaECL B Solution
  • QuickBlock™ Western Solution Kit
    • QuickBlock™ Western Blocking Solution
    • QuickBlock™ Western Antibody Dilution Buffer
    • Western Washing Buffer (10X)
    • QuickBlock™ Western Secondary Antibody Dilution Buffer
  • Extra Thick Filter Paper
  • Ultrapure Water
  • 95% Ethanol
  • Anhydrous Methanol
  • His Tag Mouse Monoclonal Antibody
  • HRP-labeled Goat Anti-Mouse IgG(H+L)

  Table 1: Primary Antibody Working Solution

  Reagents	Volume (mL)
  His Tag Mouse Monoclonal Antibody	0.005
  QuickBlock™ Western Antibody Dilution Buffer	5

  Table 2: Secondary Antibody Working Solution

  Reagents	Volume (mL)
  HRP-labeled Goat Anti-Mouse IgG(H+L)	0.010
  QuickBlock™ Western Secondary Antibody Dilution Buffer	5

  Procedure

  1. Balance

  • Measure 100 mL of BeyoWB™ Transfer Buffer (10X) and pour it into a clean graduated cylinder. Add ultrapure water to a total volume of approximately 700 mL. Add 200 mL of anhydrous ethanol or 210 mL of 95% ethanol and mix thoroughly. Adjust the final volume to 1 liter with ultrapure water in the graduated cylinder, mix well, and the 1X transfer buffer is ready for use.
  • Wash the gel with transfer buffer. Place the gel in a container, and add 15 mL transfer buffer. Put the container on a horizontal shaker at 40 rpm. Rinse for 5–10 minutes.
  • Cut one PVDF membrane and two extra thick filter paper to the same size of the gel.
  • Put the extra thick filter paper to a container with sufficient transfer buffer.

  2. Activate the PVDF membrane

  • Add approximately 10 mL anhydrous methanol to a container.
  • Use flat-tipped tweezers to hold one corner of the PVDF membrane, positioning it as horizontally as possible on the surface of the anhydrous methanol.
    Caution: Ensure the container is completely dry before adding anhydrous methanol; if not thoroughly dry, pre-rinse with a small amount of methanol.
  • The PVDF membrane will quickly become uniformly semi-transparent and gradually sink into the conditioning solution. The activation typically takes about 2–5 minutes.
  • Remove the PVDF membrane with flat-tipped tweezers and place it into the transfer buffer for a quick rinse of 5–10 seconds.
  • Put the PVDF membrane into another container with sufficient transfer buffer. Put the container on a horizontal shaker at 40 rpm for 1–2 minutes.

  3. Prepare the transfer sandwich in the transfer chamber

  • Place thick filter paper first, then add the PVDF membrane. Trim a small corner at the top right of the membrane to serve as a marker for orientation.
  • Place the gel on top, followed by another piece of filter paper.
    Caution: Removing air bubbles completely is critical for achieving good transfer results. A bubble roller can be used to gently roll out the bubbles at each step.
  • Add 1–2 mL of transfer buffer to the sandwich in the transfer chamber to ensure the filter paper is fully wetted.

  4. Transfer

  • Firmly clamp the transfer sandwich and lock the sandwich with the white slider, ensuring that the sandwich stays in place. Insert the transfer sandwich with the black side facing the black side of the transfer core assembly. Perform the transfer at a constant current of 25V, 1A for 11 minutes at room temperature, without cooling needed.

  5. Membrane Blocking

  • After the transfer is finished, immediately place the membrane into a washing box pre-filled with 15 mL BeyoWB™ Western Wash Buffer. Put the container on a tilting shaker at 20 rpm. Rinse for 1–2 minutes.
  • Remove the wash buffer from the box using a pipette. Add 10 mL BeyoWB™ Blocking Buffer. Put the container on a tilting shaker at 20 rpm. Rinse for 10 minutes.

  6. Primary antibody incubation

  • Remove the Blocking Solution from the box using a pipette. Add 15 mL Primary Antibody Working Solution. Put the container on a tilting shaker at 20 rpm. Rinse for 15-20 minutes.
    Caution: Recover the Primary Antibody Working Solution for future reuse. Recovered antibodies can typically be reused 5–10 times. Store diluted antibodies, including previously used diluted antibodies, at 4°C.
  • Wash the gel with wash buffer 3 times. Each time using 15 mL wash buffer, 3 minutes.

  7. Secondary antibody incubation

  • Add 15 mL Secondary Antibody Working Solution. Put the container on a tilting shaker at 20 rpm. Rinse for 15-20 minutes.
  • Wash the gel with wash buffer 3 times. Each time using 15 mL wash buffer, 3 minutes.
    Caution: Recover the Secondary Antibody Working Solution for future reuse. Recovered antibodies can typically be reused 5–10 times. Store diluted antibodies, including previously used diluted antibodies, at 4°C.

  8. Protein detection

  • Prepare the BeyoECL Working Solution. Each PVDF membrane needs 1.2 mL working solution. Mix 0.6ml of BeyoECL A solution with 0.6ml of BeyoECL B solution to make 1.2 mL of the BeyoECL Working Solution. It is recommended to use it immediately after preparation.

  • Lay a clean plastic wrap on the benchtop. Discard the remaining wash solution in the container, then place the membrane flat on the plastic wrap. Ensure there is no obvious residual wash solution on the membrane surface. If any residue is present, carefully absorb it with lens paper, taking care not to touch the membrane.

  • Add 1.2 mL of the BeyoECL Star Working Solution onto the membrane, ensuring even coverage, and let it sit for 5 minutes.

  • Discard the Working Solution and gently remove excess liquid with absorbent paper. Then proceed with chemiluminescence imaging.

    ​

Micromotor Construction & Estimation

• Gel Encapsulation Experiment

  Materials

  • 1× PBS (pH 6.8)
  • 1.5% (w/v) Sodium alginate solution
  • Nano calcium carbonate
  • Liquid paraffin containing 0.5% Span 80^®
  • Glacial acetic acid (sterile)
  • Acetate buffer (pH ≈ 5.5)
  • Saccharomyces boulardii culture at logarithmic growth phase

  Procedure

  • Centrifuge 50 mL of logarithmic-phase Saccharomyces boulardii culture (1100 × g, 5 min) to collect cells. Wash twice with deionized water, then resuspend in just enough deionized water. Measure OD value and adjust the final OD₆₀₀ to 1.946.
  • In a sterile 100 mL beaker, add 10 mL sodium alginate solution. Start the digital overhead stirrer at 310 rpm.
  • Mix 1 mL yeast suspension with 10 mL sodium alginate solution to obtain the yeast-containing sodium alginate mixture.
  • Add 0.1 g nano calcium carbonate into the sodium alginate–yeast solution and stir until well dispersed.
  • Add 55 mL liquid paraffin containing 0.5% Span 80 to achieve an aqueous-to-oil phase ratio of 1:5.
  • After emulsifying for 15 min, use a plastic dropper to add 1.38 mL glacial acetic acid dropwise within 30 min, while continuing stirring. From the start of acid addition, continue stirring for 60 min to promote calcium carbonate dissolution and initiate gelation.
  • Washing: Centrifuge at 1100 rpm for 5 min, remove the upper oil phase after microsphere sedimentation. Wash with acetate buffer (optionally containing 1% Tween 80) and centrifuge again. Repeat until no visible oil phase remains.

• Gel Degradation Test

  Materials

  • Calcium alginate hydrogel microspheres (prepared as described above)

  • Phosphate-buffered saline (PBS, pH 7.4)

  • Acetate buffer (pH 5.0)

  • 1M HCl solution （pH 1.0）

  • Deionized water

  • 1.5 mL Eppendorf (EP) tubes, labeled 1–9

  • Optical microscope

  • Vortex mixer

  Procedure

  • Sample Preparation: Collect equal volumes (e.g., 500 μL) of calcium alginate hydrogel microspheres and transfer them into nine 1.5 mL EP tubes labeled 1–9.
  • Buffer Treatment: Add 1 mL of different buffer solutions to each tube according to the following setup:
    • Tube 1–3: PBS (pH 7.4)
    • Tube 4–6: Acetate buffer (pH 5.0)
    • Tube 7–9: 1M HCl solution (pH 1.0)
  • Mix gently using a vortex mixer to ensure even suspension.
  • Incubate all tubes at 37 °C under static conditions. Record time points at 0 h, 1 h, 3 h, 6 h, 12 h, and 24 h.
  • At each time point, visually inspect and photograph the supernatant turbidity as an indicator of gel disintegration.
  • Microscopic Examination: Take a small aliquot (≈10 μL) from each tube at each time point and observe under an optical microscope (bright field) to assess changes in microsphere integrity and morphology.

• Scanning Electron Microscopy (SEM) Observation

  Materials

  • Concentrated and purified sodium alginate hydrogel
  • Liquid nitrogen

  Experimental Procedure

  • Pipette a small amount of hydrogel solution onto a copper freezing sample holder.
  • Place the holder onto the manipulator rod and insert it into the sample pretreatment chamber. Add purified liquid nitrogen (filtered with filter paper) while simultaneously applying vacuum (to form supercooled liquid nitrogen).
  • Transfer the sample into the transfer chamber. Once the vacuum levels of the transfer chamber and the sample preparation chamber are consistent, open the intermediate valve and push the transfer rod to move the sample into the preparation chamber (at −145 °C).
  • Use the blade to cut the sample.
  • Maintain the sample at −90 °C for 5 minutes to allow ice sublimation.
  • Perform ion sputtering to coat the sample with a ~3 nm gold layer.
  • Place the sample into the environmental scanning electron microscope (ESEM) for observation.

• Fabrication of Gel Micromotors Using a Microfluidic Chip

  Materials

  • 2% Sodium Alginate Solution
  • 10% Sodium Carbonate Solution
  • 1% CaCl2 Solution
  • Soybean oil containing 1% Span 80

  Procedure

  Here is a photo

  • Contents of Inlets

    • Inlet 1: Soybean oil.
    • Inlet 2: 2% sodium alginate solution.
    • Inlet 3: Sodium carbonate solution with concentrations of 10%.
    • Inlet 4: 1% CaCl₂ solution.
    • Inlet 5: Soybean oil.

  • Place on a glass slide and observe morphology under an optical microscope at 10–40× magnification.

  • Flow rates:

    • Inlets 1 and 5: 70–350 μL/h
    • Inlets 2 and 3: 5–25 μL/h
    • Inlet 4: frequency synchronized with droplet cutting rate in the intersection.

• Micromotor Preparation via Macroscale Emulation

  Materials

  • 2% Sodium Alginate Solution
  • Nano calcium carbonate
  • crystal violet stain
  • 1% CaCl₂ Solution

  Procedure

  • Disperse 5 g of nano-calcium carbonate in 50 mL of sodium alginate solution.
  • Load 2% sodium alginate solution and sodium alginate solution containing 10 wt% nano-calcium carbonate into two 1 mL syringes, respectively, and connect the syringe needles to two microfluidic tubes.
  • Add 1% calcium chloride solution into a 50 mL beaker.
  • Use glue to align and fix the two microfluidic tubes, inserting their outlets into the mouth of the beaker.
  • Push the syringes to extrude the two liquids simultaneously, forming Janus droplets, which fall into the calcium chloride solution.

• Micromotor Propulsion Test in Simulated Gastric Acid Environment

  Materials

  • 1 M HCl solution
  • Gel micromotors

  Procedure

  • Add 400 mL of 1 M HCl solution into a 1000 mL beaker.
  • Place the hydrogel microspheres inside, record a video, and track their motion.

Assays

• BCA Protein Assay

  Materials

  • BCA Protein Assay Kit
    • BCA Reagent A
    • BCA Reagent B
    • Protein Standard(BSA)
    • BSA Preparation Solution
  • 1×PBS (pH = 7.4)

  Procedure

  1. Preparation of protein standard samples

  1. Add 1.2 ml of the BSA Preparation Solution to a tube containing 30 mg of Protein Standard.

  2. Take an appropriate amount of the 25 mg/ml Protein Standard and dilute it with 1×PBS to reach a final concentration of 0.5 mg/ml.

  2. Configuration of BCA working solution

  1. Based on the sample quantity, prepare an appropriate amount of BCA working solution by adding 50 volumes of BCA Reagent A to 1 volume of BCA Reagent B, and mix thoroughly.

  Table 1: BCA working solution.

  Component	Volume
  BCA Reagent A	5mL
  BCA Reagent B	100μL

  Caution: The BCA working solution remains stable at room temperature within 24 hours.

  3. Determination of protein concentration

  1. Add the Protein Standard and 1×PBS to the wells of the 96-well plate according to the quantities specified in the Table 2 below.

  Table 2: Gradient concentration Protein Standard.

  Volume of Protein Standard	Volume of 1×PBS	Final Concentration
  0 μL	20 μL	0 mg/mL
  1 μL	19 μL	0.025 mg/mL
  2 μL	18 μL	0.05 mg/mL
  4 μL	16 μL	0.1 mg/mL
  8 μL	12 μL	0.2 mg/mL
  12 μL	8 μL	0.3 mg/mL
  16 μL	4 μL	0.4 mg/mL
  20 μL	0 μL	0.5 mg/mL

  2. Add an appropriate volume of the sample to the sample wells of the 96-well plate. If the sample volume is less than 20 µl, add the standard substance diluent to make up to 20 µl.

  3. Add 200 µl of BCA working solution to each well, and incubate at 37°C for 20-30 minutes.
     Caution: When using the BCA method to determine protein concentration, the color will deepen continuously over time. And the color development reaction will accelerate due to the increase in temperature.

  4. Measure the absorbance at A₅₆₂ using an enzyme detector.

  5. Based on the corrected standard substance concentration and absorbance in the blank, plot a standard curve.

  6. Calculate the protein concentration of the sample based on the standard curve and the volume of the sample used.

• BeyoBCA Rapid Protein Assay

  Materials

  • BeyoBCA Rapid Protein Assay Kit
    • BCA Reagent A
    • BCA Reagent B
    • BCA Reagent C
    • Protein Standard (BSA)
    • BSA Preparation Solution
  • 1×PBS (pH = 7.4)

  Procedure

  1. Preparation of protein standard samples

  1. Add 1.2 ml of the BSA Preparation Solution to a tube containing 30 mg of Protein Standard.

  2. Take an appropriate amount of the 25mg/ml Protein Standard and dilute it with 1× PBS to reach a final concentration of 10mg/ml.

  2. Configuration of BCA working solution

  1. Based on the sample quantity, prepare an appropriate amount of BCA working solution, and mix thoroughly. The specific preparation method can be found in the Table 1 below.

  Table 1: BCA working solution.

  Component	Volume
  BCA Reagent A	100μL
  BCA Reagent B	96μL
  BCA Reagent C	4μL

  Caution: The BCA working solution remains stable at room temperature within 30 minutes.

  3. Determination of protein concentration

  1. Add the Protein Standard and 1×PBS to the wells of the 96-well plate according to the quantities specified in the Table 2 below.

  Table 2: Gradient concentration Protein Standard.

  Vial Number	Volume of Protein Standard	Volume of 1×PBS	Final Concentration
  A	70 μL Protein Standard(10mg/ml)	0 μL	10 mg/mL
  B	35 μL of Vial A	35 μL	5 mg/mL
  C	35 μL of Vial B	35 μL	2 mg/mL
  D	35 μL of Vial C	35 μL	1 mg/mL
  E	35 μL of Vial D	35 μL	0.5 mg/mL
  F	35 μL of Vial E	35 μL	0.25 mg/mL
  G	35 μL of Vial F	35 μL	0.125 mg/mL
  H	0 μL	35 μL	0 mg/mL

  2. Add an appropriate volume of the sample to the sample wells of the 96-well plate. If the sample volume is less than 10 µl, add the standard substance diluent to make up to 10 µl.

  3. Add 200 µl of BCA working solution to each well, mix well and leave at room temperature (around 25°C) for 5 minutes.

  4. Measure the absorbance at A₄₈₀ using an enzyme detector.

  5. Based on the corrected standard substance concentration and absorbance in the blank, plot a standard curve.

  6. Calculate the protein concentration of the sample based on the standard curve and the volume of the sample used.

• Cell Counting Assay

  Procedure

  1. Absorption spectrophotometry method

  1. Preheat the spectrophotometer 40-60 minutes in advance.

  2. Modify the wavelength to 600 nm.

  3. In Mode T, do calibration to 0 at the black cuvette and 100 at the cuvette with the solvent.

  4. Switch to Mode A and measure the OD value of the sample culture.

  5. If the OD value is significantly higher than 0.8, dilute the culture medium appropriately to a range of 0.2 to 0.8 (dilution factor D) and re-determine the value.

  2. BioTech^® Epoch 2 Instrument measurement method

  1. Preheat the enzyme label reader 2-3 minutes in advance, and set the protocol and relevant parameters.

  2. Set up two sets of duplicate culture media and the test solution, and add 200 μL to each of the 96-well plates.

  3. If the OD₆₀₀ value of the sample to be tested exceeds 1.0, then set a suitable dilution gradient of the appropriate concentration.

  3. Blood cell counting plate measurement method

  1. Dilute the yeast culture solution by $d$ ($d = 1$ if not diluted)

  2. Add 10 μL drop to the area between the cover glass and the inclined surface of the counting plate to allow the culture medium to seep in and fill the counting chamber completely at one time.

  3. Count the number of cells in the yellow squares in the microscopic count chart, and then calculate the average value $m$.

  4. The number of cells in each yellow square should be controlled within 50 to 100 (2 to 3 adhered cells are counted as 1 cell; if more than 4 to 5 adhered cells are present, the yeast solution needs to be diluted again before counting). The range of variation should not exceed 15; otherwise, the cells are considered dispersed unevenly and the solution needs to be re-added for counting.

     

     • Cell density in diluent: $n\mathrm{(cells/mL)}=m×25×10^4$

     • Original cell density: $N\mathrm{(cells/mL)}=d·n$

• Florescence Reporting

  Materials

  • PBS
  • Protein Samples
  • 96-well plate

  Procedure

  1. Add 200 μL of each protein sample to the 96-well plate, and add 200 μL of PBS to another well as a control.
  2. Detect the samples using the microplate reader’s default wavelengths (excitation wavelength 488 nm, emission wavelength 528 nm).
  3. Use the fluorescence intensity data provided by the microplate reader to determine the EGFP content in the protein samples, thereby inferring the expression of the exogenous plasmid in the cells.

• Genotype Identification

  Materials

  • Yeast Colony PCR Kit with Alkaline Lysis
    • Yeast Lysis Buffer
    • Neutralization Buffer
    • Yeast Colony PCR Mix (Green, 2X)
  • 1M sobitol
  • 100mM EDTA
  • 14nM β-mercaptoethanol
  • PCR Master Mix
  • Agarose
  • 1×TAE
  • DNA Marker
  • M5 GelRed Plus nucleic acid dye (10000×)

  Procedure

  1-1. Yeast Colony PCR

  1. Add 10 µl of Yeast Lysis Buffer to each tube. Use a sterile pipette tip to pick a yeast monoclonal colony (0.2-1 mm in size) and transfer it into a PCR tube containing 10 µl of Yeast Lysis Buffer. Mix by pipetting or vortexing, and centrifuge briefly to collect the liquid at the bottom of the tube. Simultaneously, label the colony on the plate.

  2. Heat the tube at 95°C for 5 minutes in a PCR machine to fully release the yeast DNA. Then add 10 µl of Neutralization Buffer and mix thoroughly.

  3. Prepare the PCR mix on ice, and then start PCR with the steps below.

     Table: PCR System Composition

     Reagent	Volume
     Ultrapure Water	7.4 μL
     primer mix	2×0.8 μL
     Yeast Colony PCR mix	10 μL
     template	1 μL
     total	20 μL

     Table: PCR Process Overview

     Steps	Temperature	Time
     Initial Denaturation	94°C	3min
     Denaturation	94°C	30s
     Annealing	55°C	30s
     Extension	72°C	1min/kb
     Go to STEP2 for 30 cycles
     final Extension	72°C	10min
     hold	4°C

  1-2. Genome Extraction & PCR

  Table: Yeast Lysis Buffer

  Component	Concentration
  Sorbitol	1M
  EDTA	100nM
  β-mercaptoethanol	14nM

  1. Centrifuge to collect yeast pellet. Resuspend in 600 μL of yeast lysis buffer, add 200 U of lyticase, and incubate at 30°C for 30 minutes. Centrifuge at 300g for 10 minutes to collect the precipitate, and discard the supernatant. Resuspend the precipitate in 180μL of Sample Lysis Buffer A.

  2. Add 20 μL of Proteinase K and vortex to mix. Incubate in a 55°C water bath until complete lysis is achieved.

  3. Vortex vigorously at maximum speed for 15s. Add 200 μL of Sample Lysis Buffer B and vortex to mix. Incubate at 70°C for 10 minutes. Add 200 μL of absolute ethanol and vortex to mix.

  4. Transfer the mixture from step e to a DNA purification column. Centrifuge at 6000g for 1min. Discard the flow-through in the collection tube.

  5. Add 500 μL of Wash Buffer I and centrifuge at 6000g for 1min. Discard the flow-through in the collection tube.

  6. Add 600 μL of Wash Buffer II and centrifuge at 12,000rpm for 1 min. Discard the flow-through in the collection tube.

  7. Centrifuge again at 12,000rpm 1 min to remove residual ethanol.

  8. Place the DNA purification column into a clean 1.5 mL microcentrifuge tube. Add 30μL of Elution Buffer and let it stand at room temperature for 1–3 min. Centrifuge at 12,000 rpm for 1 minute. The resulting liquid is the purified total DNA. Use the Nanodrop to measure the concentration of total DNA.

  9. Prepare the PCR mix, and then start PCR with the steps below

     Table: PCR System Composition

     Component	Volume
     Water	(21-x) μL
     Template	x μL (1μg)
     Primer Mix	2×2 μL
     PCR Master Mix	25 μL
     total	50 μL

     Table: PCR Process Overview

     Steps	Temperature	Time
     Initial Denaturation	92°C	3min
     Denaturation	92°C	30s
     Annealing	55°C	30s
     Extension	68°C	1min/kb
     Go to STEP2 for 30 cycles
     final Extension	68°C	10min
     hold	4°C

  2. Agarose Gel Electrophoresis

  1. Weigh 0.16 g of agarose and dissolve it in 1× TAE buffer to a total volume of 20 mL. Heat the mixture in a microwave until completely dissolved. Add 2 μL of M5 GelRed Plus nucleic acid dye (10,000×), mix gently, and pour the solution into a medium-sized gel tray. Insert a comb and allow the gel to solidify at room temperature.
  2. During loading, add DNA marker into one well on the side of the gel. Run electrophoresis at 150 V for approximately 30 min. Observe the electrophoresis results using a gel imaging system and determine the cell genotype based on the band pattern.

• Modified Immunocytochemistry (ICC) Assay

  Materials

  • PLL storage solution(10X)
  • 0.2 μm membrane filter
  • Immunol Staining Wash Buffer
  • Immunol Staining Fix Solution
  • QuickBlock™ Blocking Buffer for Immunol Staining
  • QuickBlock™ Primary Antibody Dilution Buffer for Immunol Staining
  • QuickBlock™ Secondary Antibody Dilution Buffer for Immunofluorescence
  • Primary antibody
  • Secondary antibody
  • Calcofluor White (5mM, 200X)

  Table: PLL storage solution(10X)

  Component	Amount
  poly-L-lysine	5mg
  ddH$_2$O	5ml

  Caution: Dissolve compounds thoroughly. Store at -20℃.

  Procedure

  1. Equipment Preparation

  1. Prepare the PLL slides: In the petri dish, immerse the new clean slides in the PLL working solution at room temperature for 5 minutes (1-2 mL per slide), then dry them in a 60℃ oven for 1 hour and leave them at room temperature overnight for drying.

     Caution: If necessary, clean the slides with 70% ethanol containing 1% HCl.

  2. Sample Preparation

  1. Dilute the PLL stock solution 10 times and filter sterilize it through a 0.2 μm membrane filter to obtain the PLL working solution.

     Caution: Use immediately after preparation. Store it at 4℃ for a short period.

  2. Cultivate 5 ml of yeast expressing protein A to the early stage of logarithmic growth. (10$^6$~10^7^ cells/ml)

  3. Centrifuge at 5000×g for 1 minute, then discard the supernatant. Wash with 1×PBS (pH=7.4).

  4. Repeat step 2 twice.

  5. Add 1 mL of cold immunostaining fixative, and fix at 4℃ for 10 minutes or longer.

  6. Centrifuge at 5000×g for 1 minute, then discard the supernatant. Wash with the Immunol Staining Wash Buffer for 5 minutes.

  7. Repeat step 5 twice.

  8. Centrifuge at 5000×g for 1 minute. Discard most of the supernatant. Keep approximately 50 μl. Place it on a PLL slide and let it air dry slightly until the cells do not move randomly.

  3. Blocking

  1. Add an appropriate amount (about 1-2 mL per slide) of QuickBlock™ Blocking Buffer for Immunol Staining to immerse the slide.
  2. Slowly shake it on a side-shaking incubator for 15 minutes.

  Caution: From the beginning of the blocking process, it is essential to ensure that the samples are kept moist to prevent them from drying out. Otherwise, a very high background level will be easily generated.

  4. Protein B Incubation

  1. Use the QuickBlock™ Primary Antibody Dilution Buffer for Immunol Staining to dilute the protein B, obtaining the protein B working solution. Approximately 1 mL is needed for each sample.

  2. Remove the sealing solution from slides and carefully absorb the remaining liquid along the edges using absorbent paper. Immediately add the protein B working solution, and gently shake the mixture at room temperature or 4°C on a rotator for 1 hour.

     Caution: If the result is not satisfactory, the sample can be gently shaken at 4℃ and incubated overnight.

  3. Add the Immunol Staining Wash Buffer, and gently shake it on the side-shaking incubator for 5 minutes. After removing the wash buffer, add new wash buffer again and shake for another 5 minutes. It should be washed twice.

     Caution: If the background color is still too high, appropriately extend the washing time and increase the number of washes.

  5. Primary Antibody Incubation

  1. Dilute the primary antibody with the QuickBlock™ Primary Antibody Dilution Buffer to obtain the primary antibody working solution. Approximately 1 mL of the working solution is needed for each slide.

  2. Remove the sealing solution from slides and carefully absorb the remaining liquid along the edges using absorbent paper. Immediately add the primary antibody working solution, and gently shake the mixture at room temperature or 4°C on a rotator for 1 hour.

     Caution: If the result is not satisfactory, the sample can be gently shaken at 4℃ and incubated overnight.

  3. Recycle the primary antibody.

  4. Add the Immunol Staining Wash Buffer, and gently shake it on the side-shaking incubator for 5 minutes. After removing the wash buffer, add new wash buffer again and shake for another 5 minutes. It should be washed for 3 times.

     Caution: If the background color is still too high, appropriately extend the washing time and increase the number of washes.

  6. Secondary Antibody Incubation

  1. Dilute the secondary antibody with the QuickBlock™ Secondary Antibody Dilution Buffer for Immunofluorescence. Adjust the dilution ratio appropriately based on the intensity of the fluorescence. Approximately 1 mL of the working solution is needed for each sample.

  2. Discard the washing solution and carefully absorb the remaining liquid with absorbent paper along the edges. Immediately add the diluted secondary antibody working solution, and gently shake it at a side swing shaker at room temperature or 4°C for 1 hour.

  3. Recycle the secondary antibody.

  4. Add the immunostaining Staining Wash Buffer, and gently shake it on the side-shaking incubator for 5 minutes. Keep in dark place. After removing the wash buffer, add new wash buffer again and shake for another 5 minutes. It should be washed for 3 times.

     Caution: If the background color is still too high, appropriately extend the washing time and increase the number of washes.

  7. Calcofluor White counterstaining

  1. Dilute the Calcofluor White stock solution with ddH$_2$O at a ratio of 1:200. According to the intensity of the fluorescence, the dilution ratio can be appropriately increased or decreased. Approximately 1 mL of working solution is needed for each piece.
  2. Discard the washing solution and carefully blot the edges with absorbent paper to dry them. Then, immediately add the diluted CFW staining solution and perform the staining process for 10 minutes.
  3. (Optional) Add the immunostaining washing solution, then gently shake it on a rotator table. Keep in dark place for 2 to 3 minutes for washing, and then remove all the washing solution.

  8. Fluorescence Detection

  1. Drop a drop of anti-fluorescence quenching mounting solution onto the slide, then cover it.

     Caution: When covering the slide, try to avoid bubbles. Do not allow the slide to move relative to the sample.

  2. Observe under a fluorescence microscope.

  3. If preservation is required, cover the specimen with paraffin or nail polish and store it at -20℃.

• Modified Co-Immunoprecipitation (Co-IP) Assay

  Materials

  • Extracted HopQ protein

  • BeyoMag™ His-tagged protein purification agarose magnetic beads(IDA-Ni)

  • Ultra pure water

  • 1xTBS

    Table: The components of 1×TBS (PH=7.25-7.55) .

    Concentration	Component
    50 mmol/L	Tris-HCl
    150 mmol/L	NaCl

  • Elution Buffer

    Table: The components of Elution Buffer.

    Concentration	Component
    10 mmol/L	Tris-HCl
    500 mmol/L	NaCl

    • Filter with a 0.22 μm filter membrane and store at 4℃ for future use.

  • Binding/Wash Buffer

    Table: The components of Binding/Wash Buffer.

    concentration	Component
    10 mmol/L	Tris-HCl
    500 mmol/L	NaCl
    500 mmol/L	Imidazole

    • Filter with a 0.22 μm filter membrane and store at 4℃ for future use.

  • 20% Ethanol

  • 5mg/ml c-Myc Peptide solution

  • Primary Antibody: His Tag Mouse Monoclonal Antibody

  • Secondary Antibody: Garlic peroxidase-labeled goat anti-mouse IgG(H+L)

  Procedure

  1. Purify HopQ Protein

  1. Preparation of BeyoMag™ His-tagged Protein Purification Agarose Magnetic Beads (referred to as IDA-Ni Magnetic Beads)
     • Gently resuspend the IDA-Ni magnetic beads with a pipette.
     • Calculate the amount of magnetic bead suspension needed based on 1 mL of suspension for every 5-10 mg of target protein (with a molecular weight of approximately 60 kD). Transfer an appropriate amount of IDA-Ni magnetic beads to a clean centrifuge tube, and remove the supernatant by magnetic separation.
     • Add an equal volume or a slightly larger volume of Binding/Wash Buffer to wash the magnetic beads.
     • Use a pipette to gently agitate and resuspend the IDA-Ni magnetic beads. Place them on the magnetic stand (FMS012/FMS024) and let them separate for 10 seconds. Remove the supernatant. Repeat this process twice.
  2. Binding
     • Add protein to equilibrated beads and incubate at room temperature on a rotary mixer for 20 minutes.
     • Place on the magnetic rack and let it separate for 10 seconds. Remove the excess supernatant.
     • Add 2 ml of Binding/Wash Buffer and gently pipette to resuspend the IDA-Ni magnetic beads.
     • Place on the magnetic rack and let it separate for 10 seconds. Remove the supernatant. Repeat this washing process three times.
  3. Washing
     • Add 0.5 ml of Elution Buffer, gently invert the centrifuge tube several times.
     • suspend and incubate for 5 minutes, perform magnetic separation, and collect the eluate into a new centrifuge tube.
     • Repeat the above steps once.

  2. Yeast co-culture

  1. Select a single colony of the positive yeast strain C1ND and inoculate it into 5 mL of LB medium (containing 100 μg/mL Amp) for overnight cultivation at 37°C.

  2. The overnight culture is inoculated at a ratio of 1:100 into 100 mL of LB (containing 100 μg/mL Amp), and then culture at 37°C and 200 rpm until the OD600 reach 0.5-0.6.

  3. Add IPTG to achieve a final concentration of 0.5 mM, take three 15mL centrifuge tubes, and aliquot 10 mL of the bacterial solution into each tube.

  4. Three centrifuge tubes are labeled and separately added with the purified product of HopQ. Incubate the cultures for 4-24 hours.

     Table: The handling method of Tubes.

     Label	Component
     Tube A	1mL HopQ protein
     Tube B	3mL HopQ protein
     Tube C	5mL HopQ protein

  3. Lysis

  1. Centrifuge at 12000g for 2 minutes at 4°C. Add 3 μL of protease inhibitor.
  2. Resuspend the cells with 0.3 ml of BeyoLytic™ bacterial activity protein extraction reagent. Incubate at room temperature for 15 minutes.
  3. Harvest soluble proteins by centrifugation at 12000g for 5 minutes at 4°C.

  4.Pull down

  1. Preparation of Anti-Myc Beads.
     • Blot the Anti-Myc magnetic beads and add 1X TBS until the final volume reaches approximately 0.5 mL. Place on the magnetic rack and let it separate for 10 seconds. Remove the supernatant. Repeat this process twice.
     • Using 1X TBS to resuspend the Anti-Myc magnetic beads according to the initial volume.
  2. Binding and Washing
     • Add Anti-Myc magnetic beads in a ratio of 10 μL or 20 μL per 500 μL of protein sample to the sample, and then place it on a shaking incubator or a rotary mixer.
     • Incubate at room temperature for 2 hours or at 4°C overnight.
     • Place on the magnetic rack and separate for 10 seconds. Retain a portion of the supernatant for testing the effect of immunoprecipitation, and discard the rest of the supernatant.
     • Add 500 μl of 1X TBS and gently pipette to resuspend the Anti-Myc magnetic beads. Place on a magnetic stirrer and let it separate for 10 seconds. Remove the supernatant.
     • Repeat this washing process three times.
  3. Elution
     • For every 10-20 μL of the original magnetic beads, add 100 μL of 3× c-Myc peptide elution solution (150 μg/ml), mix well, and then place it on a shaking incubator or a rotary mixer at room temperature for 30-60 minutes.
     • Place on the magnetic rack and separate for 10 seconds. Transfer the supernatant to a new centrifuge tube.
     • The eluted Myc-tagged protein should be stored at 4°C for future use, or at -20°C or -80°C for long-term preservation.

  5. analysis

  1. Perform SDS-PAGE (using 4% precast gradient gel, Bio-RAD).
  2. Analyze the resolved proteins by performing a Western Blot using His Tag Mouse Monoclonal Antibody.
  3. Use secondary antibody(Garlic peroxidase-labeled goat anti-mouse IgG) for detection.
  4. Based on the known molecular weights of HopQ and C1ND, determine the rationality of the position where the interaction bands appear.

• Pull-down Assay

  Materials

  • Protein sample

  • Anti-Myc Magnetic Beads

  • c-Myc peptide elution buffer

  Procedure

  1. Preparation of Anti-Myc Magnetic Beads

  • Gently resuspend the Anti-Myc beads using a pipette. Transfer an appropriate volume of bead suspension (10 μL or 20 μL per 500 μL of sample) into a clean microcentrifuge tube. Add 1× TBS to a final volume of approximately 0.5 mL.

  • Gently pipette up and down to completely resuspend the beads. Place the tube on a magnetic rack for 10 seconds and discard the supernatant. Repeat this wash step twice.

  • Resuspend the beads in 1× TBS to the original volume.

  2. Immunoprecipitation

  • Bead incubation: Add Anti-Myc beads to the protein sample at a ratio of 10 μL or 20 μL of bead suspension per 500 μL of protein sample.

  • Incubate in a rotating mixer or nutator at room temperature for 2 hours, or overnight at 4°C.
    Note: Bead aggregation or sheet-like appearance during incubation is normal and does not affect the result.

  • Magnetic separation: After incubation, place the tube on a magnetic rack for 10 s. Save a portion of the supernatant to assess immunoprecipitation efficiency, discard the rest.

  • Wash: Add 500 μl of 1× TBS and gently resuspend the beads by pipetting. Place on the magnetic rack for 10 s and discard the supernatant. Repeat this wash step three times.

  3. Elution

  • Preparation of c-Myc peptide elution buffer solution: Mix 30 μL of 5 mg/mL c-Myc peptide solution with 970 μL of sterile water to obtain 1 mL of 150 μg/mL elution buffer.

  • For every 10–20 μL of original bead volume, add 100 μL of 3× c-Myc peptide elution buffer (150 μg/mL). Mix gently and incubate in a rotating mixer or nutator at room temperature for 30–60 minutes, or at 4°C for 1–2 hours. To improve elution efficiency, extend the incubation time or perform repeated elution.

  • After incubation, place the tube on a magnetic rack for 10 s and transfer the supernatant to a new microcentrifuge tube. The supernatant contains the eluted Myc-tagged protein.

  • Store the eluted Myc-tagged protein at 4 °C for short-term use, or at −20 °C for long-term storage.