SNX-BARs are membrane remodeling proteins that play key roles in human disease. Using our methods, we can determine their precise biophysical properties to facilitate lipid binding and membrane remodeling. Purifying the SNB-BAR proteins from yeast cells allows the acquisition of native homo and hetero SNX-BAR dimers while minimizing the toxicity and insolubility frequently encountered with non-native expression systems.
Our methods can be used to understand the lipid specificities of all yeast SNX-BARs or to produce recombinant SNX-BAR proteins from any other organism. A correct extruder assembly is essential to preventing liposome loss during the extrusion process. For beginning students or those new to the field, visually observing the most critical steps will greatly improve your chances of success.
Begin by inoculating a large swab of yeast cells into a flask at least four times the volume of the culture containing 50 milliliters of standard YP medium supplemented with 2%raffinose and 0.1%glucose as a carbon source and grow the culture overnight in a 30 degree Celsius shaker. The next morning, inoculate the 50 milliliter pre-culture in one liter of standard YP medium with 2%raffinose and 0.1%glucose in a 2.8 liter volume baffled Fernbach flask for a four to five-hour culture in the shaking incubator. When the optical density at 600 nanometers reaches 0.2, add galactose to a final concentration of 2%to the cells for an overnight culture in the shaking incubator.
The next morning, harvest cells by centrifugation and transfer the yeast pellet into a 50 milliliter conical tube. Resuspend the pellet in 15 milliliters of purification buffer to achieve a final volume of about 30 milliliters and load the cell suspension onto a homogenizer chilled to four degrees Celsius. Lyse the samples at 20 to 25, 000 pounds per square inch for two to three rounds and transfer the lysate into a new 50 milliliter conical tube on ice.
Immediately clear the cell lysate by centrifugation and carefully transfer the supernatant into a new tube. Next, equilibrate 300 microliters of IgG Sepharose beads with three re-suspensions in purification buffer before adding the beads to the cleared cell lysate for a two-hour incubation with rotating at four degrees Celsius. At the end of the incubation, add the beads to a 10 milliliter chromatography column and allow the unbound lysate to flow through.
Wash the beads 10 times with one milliliter of wash buffer per wash allowing each wash to flow through completely before adding the next volume. After the last wash, use a pipette and fresh buffer to transfer the beads into a microcentrifuge tube. Bring the total volume of the beads to 500 microliters with fresh wash buffer and remove the beads from the lysate with two microliters of 10 milligram per milliliter TEV protease for an overnight incubation with rotation at four degrees Celsius.
The next morning, use a 27 gauge needle to remove the supernatant completely and assess the protein purity by 10%polyacrylamide SDS page. Then quantify the concentrated proteins using the Bradford protein assay according to standard protocols and store the protein for up to one week at four degrees Celsius. To prepare the liposomes, use glass syringes to transfer stock lipids into a clean glass culture tube to achieve a final lipid mixture of 1%phosphatidylinositol-3-phosphate, 20%ergosterol, 30%phosphatidylserine, phosphatidylcholine as outlined in the table.
Depending on the solvents each lipid is resuspended in, the mixture may turn cloudy upon the addition of each lipid. Carefully dry down the lipid mixtures by directing low-flow nitrogen gas at the mixtures in a circular motion to treat the lipids uniformly at the bottom of the glass tube. Then wrap the glass culture tube with foil leaving the opening uncovered and further dehydrate the lipids in a vacuum for one hour.
At the end of the incubation, add 400 microliters of binding buffer to each vial to completely rehydrate the lipids at a final liposome concentration of 2.5 millimolar before resuspending the lipids at medium speed on a vortex at room temperature for 30 minutes. The buffer should become cloudy as the lipids are resuspended. Transfer the resuspended liposomes to a microcentrifuge tube and freeze the tubes in liquid nitrogen followed by thawing in a 37 degree Celsius water bath seven to eight times.
In a chemical fume hood, clean two one milliliter glass syringes with a full volume of chloroform three times per syringe to remove any residual lipids and equilibrate each syringe with two volumes of ultrapure water and two volumes of binding buffer. Assemble a mini-extruder according to the manufacturer's recommendations. Then submerge two pieces of filter support in one 200 nanometer membrane in binding buffer.
It is important to assemble the mini-extruder so that it is completely airtight. And a good way to check for leaks is to pass buffer through the device. Sandwich the membrane between the filter supports and place the membrane into the mini-extruder.
Using one of the equilibrated one milliliter syringes, pass a volume of binding buffer similar to the volume of the liposome mixture through the mini-extruder and use the other syringe to drop the liposome mixture inverting the microcentrifuge tube to collect the last of the liposomes into the tube cap for their collection. Then extrude the liposomes through the 200 nanometer membrane 19 to 21 times and collect the extruded liposomes into a new microcentrifuge tube. Before performing the SNX-BAR liposome binding and tubulation assays, pre-clear the purified protein by ultracentrifugation and transfer the supernatant to a new microcentrifuge tube without disturbing the pellet.
Next, incubate four micromolar of purified SNX4-ATG20 and 2.5 millimolar of liposomes in a total reaction volume of 20 microliters for a 30-minute incubation at 30 degrees Celsius. To visualize and quantify the liposome tubulation, at the end of the incubation, immediately spot the samples onto a carbon-coated copper mesh grid and negative stain with 1%uranyl acetate. Then analyze the samples on a transmission electron microscope at 200 kilovolts.
For liposome binding and sedimentation, transfer 20 microliters of the reaction to a polycarbonate centrifuge tube and spin the sample with a compatible router in an ultracentrifuge. At the end of the centrifugation, carefully transfer the supernatant to a new microcentrifuge tube and resuspend the pellet in 40 microliters of SDS page sample buffer. Transfer the pellet suspension to a new microcentrifuge tube and add 20 microliters of sample buffer to the supernatant.
Then load equivalent sample volumes of pellet and supernatant onto a 10%polyacrylamide SDS page gel and perform Coomassie staining to visualize the SNX-BARs bound to the liposomes. To check for proper induction of SNX-BAR expression, a western blot against the tandem affinity purification tag can be used as the protein levels of the SNX-BARs may not be detected via Coomassie stain. After purification of the SNX-BAR heterodimer, the bands of the two SNX-BARs should appear in a one-to-one stoichiometric ratio and there should be little to no contaminating bands.
In this representative analysis, SNX-BAR heterodimers were expressed, purified, and bound to synthetic liposomes as demonstrated. Note that MVP1 formed homodimers and was expressed in bacteria as expected. SNX-BAR binding to varying liposome compositions could be quantified by densitometry to assess the effects of phosphatidylserine for example on liposome binding.
Electron microscopy imaging allows the measurement of SNX4-ATG20 liposome tubules with most tubules typically possessing diameters between 14 to 26 nanometers. Note that purified SNX-BARs can be reconstituted with cargo capture complexes on liposomes to understand how full assemblies can influence membrane remodeling. When comparing the binding of different combinations of purified SNX-BAR dimers to synthetic liposomes composed of different lipids, SNX-BAR proteins were found to possess distinct lipid binding preferences.
Always work in a hood when handling chloroform and be sure to wear the proper PPE when handling sharps or glass material.
