Expression, Purification, and Liposome Binding of Budding Yeast SNX-BAR Heterodimers

1. Yeast Strain Construction

1. Begin with TVY614 (pep4Δ::LEU2 prb1Δ::hisG prc1Δ::HIS3)¹¹ as the parent strain. This strain is deficient for vacuolar proteases, which contribute to the majority of protein degradation after cell lysis, and therefore allows for a cleaner and more efficient purification.
2. Design primers¹² and integrate tandem affinity purification (TAP) tag at the C-terminus of Atg20 (SNX-BAR ORF 1) using homologous recombination. Use polymerase chain reaction (PCR) to confirm integrations (Figure 2).
3. Perform a western blot of cell lysate against the TAP tag to confirm proper integration¹³.
   NOTE: We recommend harvesting 3 OD (1 OD ≈ 1 x 10⁷ cells) of cells for SDS-PAGE and western blot verification. Note that integration of the TAP tag should occur prior to replacing the endogenous promoters with the GAL1 promoter to allow for easier TAP tag verification via western blot.
4. Replace endogenous Snx4 (SNX-BAR ORF1) and Atg20 (SNX-BAR ORF 2) promoters with untagged GAL1 promoter using sequential homologous recombination and transformation steps of each individual ORF¹⁴.
5. Use flanking primers outside of the integration sites to PCR confirm successful integrations (Figure 2). This will result in a null phenotype of the targeted SNX-BARs in the absence of galactose in the growth medium.

2. Yeast Induction and SNX-BAR Dimer Purification

NOTE: Yeast cells can be propagated on standard YPD (yeast extract, peptone, and 2% glucose) agar plates as the modifications are chromosomally integrated.

1. Inoculate a large swab of cells into 50 mL of standard YP (yeast extract and peptone) medium with 2% raffinose and 0.1% glucose as carbon source in a flask at least 4x the volume of the culture and grow overnight in 30 °C shaker to allow proper aeration. Expect that growth in this medium will be slower compared to standard YPD.
2. Next morning, use the 50 mL preculture to inoculate into 1 L of standard YP medium with 2% raffinose and 0.1% glucose and grow for 4-5 h in 30 °C shaker.
   NOTE: The volume of preculture used to inoculate 1 L culture may be adjusted depending on the OD₆₀₀ of the preculture. The OD₆₀₀ of the 1 L culture after inoculation should be around 0.2 to allow for at least two doublings during the 4-5 h growth.
   1. Use a baffled Fernbach flask for growth to allow proper aeration, a 2.8 L volume flask is sufficient. Less aeration may result in slower growth and smaller cell pellet upon harvesting.
3. Check OD₆₀₀ to make sure culture is in the log phase (0.5-1) after 4-5 h of growth. Depending on the growth of the strain, make adjustments to the growth time to allow for at least two doublings. Add to 2% galactose and grow overnight in 30 °C shaker.
   NOTE: The OD₆₀₀ after the overnight growth may vary but culture should be saturated. Note that cells do not need to be removed from 0.1% glucose before addition of 2% galactose for this growth protocol. We recommend harvesting 3 OD of uninduced and induced cells at this step for SDS-PAGE and western blot for verification (Figure 3A,B, Lane 1-2).
4. Harvest cells by centrifugation at 4500 x g for 15 min. A swinging bucket rotor that accommodates the 1 L volume culture may be used here.
5. Transfer the yeast pellet into a 50 mL conical tube; a second centrifugation step may be performed as needed. The cell pellet will generally be around 10-15 mL in volume as measured by graduation markings and may be used immediately or stored at -80 °C.
   1. Resuspend pellet in 15 mL purification buffer (50 mM Tris pH 7.4, 300 mM NaCl, 1.5 mM MgCl₂, 1 mM Dithiothreitol (DTT), protease inhibitor cocktail) to make final volume around 30 mL.
6. Chill a homogenizer 4 °C before use and equilibrate it with purification buffer. Lyse cells using a mechanical cell disruptor or homogenizer. Load sample into the homogenizer and lyse at 20,000-25,000 psi for 2-3 rounds; note that as cells become lysed, more input pressure is required to maintain 20,000-25,000 psi. Collect the cell lysate in a 50 mL conical tube on ice.
   NOTE: If additional samples need to be lysed, the homogenizer should be thoroughly cleaned and equilibrated before loading the next sample. Keep all cell lysates on ice.
7. Immediately clear the cell lysate at 35,000 x g for 1 h at 4 °C. Carefully transfer the supernatant into new tube. Note that the lipids from cell lysis will float to the top during centrifugation and will not affect purification.
   NOTE: We recommend saving 0.5-1% of the lysate and pellet for SDS-PAGE samples. Typically, no major differences are observed and an additional western blot may be done to confirm TAP protein solubility (Figure 4, Lanes 1 and 2, respectively).
8. Equilibrate 300 µL of IgG sepharose beads with purification buffer. Add to the cleared cell lysate and incubate for 2 h, rotating at 4 °C.
9. Collect beads in a 10 mL chromatography column and allow unbound lysate to flow through.
10. Wash beads using 10 mL of wash buffer (50 mM Tris pH 7.4, 300 mM NaCl, 1.5 mM MgCl₂, 1 mM DTT), adding 1 mL at a time and allowing it to flow through completely.
    NOTE: We recommend saving 2% of the bound-IgG beads for SDS-PAGE sample. Typically, we observe four major bands; two SNX-BAR proteins and IgG Heavy and Light Chains (Figure 4, Lane 3). We recommend saving equivalent amounts of 'eluate' and 'IgG beads after TEV' to compare for TEV cleavage efficiency.
11. Collect beads and transfer them to a microcentrifuge tube. Add to 500 µL of total volume with fresh wash buffer and 2 µL of 10 mg/mL TEV protease and incubate overnight, rotating at 4 °C.
12. Next morning, remove the supernatant completely using a 27 G needle and assess protein purity by 10% polyacrylamide SDS-PAGE (Figure 4, Lane 4).
    NOTE: We typically obtain 500 µL of 0.5-1 mg/mL of 95% pure heterodimer (Figure 4, Lane 4). Additional purification using calmodulin resin can also be done, however we typically see a significant reduction in yield and recommend stopping here if purity is >90%. TEV does not interfere with liposome binding assays, though TEV can be additionally removed using Ni-NTA agarose beads¹⁵.
13. To concentrate, transfer the sample to a 0.5 mL centrifugal filter with 10 KDa cutoff and centrifuge according to manufacturer's instructions to 50 µL or less. Quantify the concentrated proteins using the Bradford protein assay. Store at 4 °C and use within one week.

3. Liposome Preparation

1. Purchase commercially available lipids: phosphatidylserine (PS), PI3P, ergosterol, and phosphatidylcholine (PC). If needed, resuspend in recommended solvent to make stock lipids.
   NOTE: Lipids are resuspended in a methanol/chloroform mixture per manufacturer's recommendations. Make sure lipid stocks are clear and warmed to room temperature before using. Resuspended lipids can be stored under argon gas and sealed using wax film (or equivalent) at -20 °C for 6-12 months or until loss of activity is observed.
2. Calculate the volume required of each lipid stock to create a mixture with the desired lipid composition (see Table 1). Assume a total of 1 mole of lipids in the lipid mixture.
3. Perform this step in a chemical fume hood. Clean glass syringes by drawing up a full syringe volume of chloroform and discarding it in a waste container. Repeat two more times. When transferring chloroform, use only glass syringes or pipettes. When drawing up chloroform, pull on the stopper slowly to prevent introduction of gas bubbles into the syringe.
4. Use glass syringes to transfer stock lipids as calculated into a clean glass culture tube to make a final lipid mixture of 1% PI3P, 20% ergosterol, 30% PS, PC (Table 1). Depending on the solvents each lipid is resuspended in, the mixture may turn cloudy upon addition of each lipid.
   NOTE: To vary concentrations of PS (0-30%), adjust volumes accordingly and compensate with varying PC (Table 1).
5. Carefully dry down the lipid mixture using nitrogen gas directed at the lipid mixture in a circular motion to dry lipids uniformly. Use low gas flow to keep the lipids at the bottom of the glass tube during the drying process. Wrap the glass culture tube with foil, leaving the opening uncovered, and further dehydrate in vacuum for 1 h.
6. Add 400 µL of binding buffer (50 mM Tris pH 7.4, 300 mM NaCl, 1 mM MgCl₂) to completely dehydrate lipids to make a final liposome concentration of 2.5 mM.
   NOTE: Final liposome concentration can be adjusted by adding more or less binding buffer. For instance, 200 µL binding buffer may be added to produce a liposome concentration of 5 mM if needed (see below).
   1. Resuspend lipids by shaking on medium speed on a vortex at room temperature for 30 min. Buffer should appear cloudy as lipids are resuspended.
7. Transfer resuspended liposomes to a microcentrifuge tube. From this point on, plastic pipette tips may be used as lipids are no longer resuspended in chloroform. Note that the liposome solution should look cloudy.
8. Freeze-thaw liposomes seven to eight times by submerging the microcentrifuge tube first in liquid nitrogen, then in a 37 °C water bath. The liposome mixture should appear completely frozen and solid by eye before thawing.
9. Perform steps involving chloroform in a chemical fume hood. Clean two 1 mL glass syringes by drawing up and discarding full syringe volumes of chloroform, three times each, to remove any residual lipids. Equilibrate each glass syringe with ultrapure water by drawing up two syringe volumes, then equilibrate with binding buffer by drawing up two syringe volumes.
10. Assemble the mini-extruder according to manufacturer's recommendations. Equilibrate one 200 nm membrane and two pieces of filter supports (see Table of Materials) by submerging each in binding buffer.
11. Sandwich the membrane between the filter supports and place in the mini-extruder. To reduce the dead volume in the assembled mini-extruder and to make sure the assembly is air-tight, pass a volume of binding buffer comparable to the volume of the liposome mixture through mini-extruder using the 1 mL glass syringes.
12. Use one of the 1 mL glass syringes and draw up the liposome mixture. Invert the microcentrifuge tube to collect the last of the liposome mixture in the tube cap for drawing up into the glass syringe.
13. Extrude liposomes by passing through the 200 nm membrane 19-21 times. Collect extruded liposomes in new a microcentrifuge tube.
    NOTE: Extruded liposomes should appear less cloudy than liposomes before extrusion. Liposomes should be used the same day and stored on ice. The last extrusion should place liposomes in the syringe opposite of the one it began in.

4. SNX-BAR Liposome Binding and Tubulation

1. Preclear purified protein at 100,000 x g in an ultracentrifuge for 20 min at 4 °C prior to conducting liposome binding and sedimentation experiments. Remove the supernatant and transfer it to a new microcentrifuge tube; do not disturb the pellet if there is one.
2. To perform liposome binding and tubulation assays, incubate 4 µM purified Snx4-Atg20 and 2.5 mM liposomes in a total reaction volume of 20 µL, varying the volume of liposomes added.
   NOTE: In the same experiment, the same volume of liposomes should be used.
3. Incubate the reaction at 30 °C for 30 min.
   NOTE: We suggest maximizing the amount of 2.5 mM liposomes added to each reaction by using no less than 10 µL liposomes to allow for visualization during sedimentation. If 10 µL of 2.5 mM liposomes is used in a 20 µL reaction, the final concentration of liposomes will be 1.25 mM. If the purified proteins are dilute and more volume is required for 4 µM protein, the lipids may be resuspended in 200 µL during the rehydration step to double the concentration of liposomes (see Step 3.6); however, this will require knowing the protein concentration prior to making the liposomes.
4. Visualize and quantify liposome tubulation.
   1. Process liposome binding reactions immediately for electron microscopy analysis. Spot samples onto a carbon-coated copper mesh grid and negative stain using 1% uranyl acetate (Figure 5B)¹⁶.
   2. Analyze samples on a transmission electron microscope (200 kV).
   3. Use image analysis software to measure and quantify tubule diameter. To accurately quantify tubule diameter of a single tubule, take three diameter measurements along the length of a tubule and average (Figure 5B).
      NOTE: Two-way analysis of variance was used to determine statistical significance (Figure 5E). Uranyl acetate is both radioactive and toxic. Proper lab safety certification is required to perform this step.
5. Liposome binding and sedimentation.
   1. Transfer the reaction (20 µL, from step 4.3) to a polycarbonate centrifuge tube and use a compatible rotor to spin at 100,000 x g in an ultracentrifuge for 20 min at 4 °C. Carefully remove the supernatant and transfer to new microcentrifuge tube. Note that the pellet should stay intact.
   2. Resuspend the pellet in SDS-PAGE 40 µL of sample buffer and transfer to a new microcentrifuge tube. Add 20 µL of sample buffer to the supernatant. Load equivalent amounts of pellet and supernatant in a 10% polyacrylamide SDS-PAGE gel and perform Coomassie staining to visualize SNX-BARs bound to liposomes (Figure 5A).
   3. To quantify the amount of SNX-BAR complex in the pellet fraction, quantify band intensities using densitometry and quantify the proportion of SNX-BAR proteins in the pellet fraction.
      NOTE: Two-way analysis of variance was used to determine statistical significance (Figure 5C).