Analysis of Neutral Lipid Synthesis in Saccharomyces cerevisiae by Metabolic Labeling and Thin Layer Chromatography

1. Growth and labeling of yeast cells with ¹⁴C-acetic acid

1. Inoculate a yeast culture by picking a colony from a plate and dispensing it into 20 mL of synthetic complete (SC) media containing 2% dextrose (see Supplementary File for the recipe of SC media). Incubate at 30 ˚C for overnight with shaking at 200 rpm.
   NOTE: Growth condition, sample volume, and treatment will differ based on the lipid(s) of interest. Prior to running full experiments, optimal growth conditions and culture volumes should be empirically determined. This protocol discusses radiolabeling of yeast cultures grown to stationary phase, a growth phase when bio-membrane and cell growth slows, and NL synthesis is very active.
2. Measure the OD₆₀₀ of the overnight culture using a spectrophotometer and dilute the yeast cell culture to a final OD₆₀₀ of 0.2 in 50 mL of fresh SC media containing 2% dextrose. Grow the cells for 24 h, or until they have reached the stationary phase (which is commonly defined by a flat lining of the cell doubling OD₆₀₀ measurement).
3. Before collecting the cells, make the quenching buffer (see Supplementary File for details). Make two 20 mL aliquots of quenching buffer for each sample (i.e., 40 mL quenching buffer for each sample) and split evenly into two 50 mL conical tubes). Store the quenching buffer aliquots at -80 °C for future use.
4. Once the cultures have reached the desired OD or growth phase, collect the cells by centrifuging at 4,100 x g for 10 min. While samples are in the centrifuge, prepare radiolabeling media by adding [1-¹⁴C] acetic acid sodium salt to dextrose-free SC media at a final concentration of 10 μCi/mL.
   CAUTION: Proper personal protective equipment (PPE) should be worn all times when working with radioactive materials. Always follow local guidelines for the proper storage, usage, and disposal of radioactive materials.
   NOTE: Both the concentration of ¹⁴C-acetic acid in the labeling media and the radiolabeling incubation time should be adjusted according to the metabolite(s) of interest. Here, a 20 min radiolabeling pulse incubation is used, which is sufficient to label NL species with a range of abundance.
5. Remove the supernatant from the pelleted cells, and wash the cell pellet once with 20 mL of dextrose-free SC media by resuspending the pellet with a pipette. Collect the cells again by centrifuging at 4,100 x g for 5 min.
6. Resuspend the cells in 1 mL of dextrose-free SC media, and transfer the cells to a labeled 2 mL microcentrifuge tube. Collect the cells again by centrifuging at 4,100 x g for 2 min.
7. Resuspend the cells once more in 500 μL of dextrose-free SC media. Pre-cool a centrifuge equipped for 50 mL conical tubes to -10 °C or on the lowest temperature setting.
8. Begin the radiolabeling period by quickly adding 500 μL of radiolabeling media to each 500 μL of cell suspension (final ¹⁴C-acetic acid concentration = 5 μCi/mL). Incubate the tubes in a rotating incubator at 30 °C for 20 min. 2 min before the end of the labeling period, transfer one 20 mL aliquot of quenching buffer for each sample from the -80 °C freezer to a bucket of ice
9. Once the radiolabeling period has ended, use a pipette to plunge the entire 1 mL sample into 20 mL of cold quenching buffer. Vortex the conical tubes for 5-10 s to ensure that the sample has been thoroughly mixed with the quenching buffer. Incubate the samples in quenching buffer for 2 min on ice.
10. Collect the cell pellet by spinning in a centrifuge at 5,000 x g for 3 min set to -10 °C or on the lowest temperature setting. While sample tubes are spinning, transfer another set of quenching buffer aliquot from the -80 °C freezer to a bucket filled with ice (i.e., one 20 mL tube of quenching buffer per sample).
11. Remove the quenching buffer supernatant from cell pellets and replace it with 20 mL fresh, cold, quenching buffer. Vortex and shake the samples until the pellet has been dislodged from the bottom of the conical tube and resuspended fully in quenching buffer. Centrifuge the samples again at 5,000 x g for 3 min at -10 °C to collect the cells.
12. Once the cells are pelleted, thoroughly remove all quenching buffer from the samples by pouring off the supernatant and removing the excess with a pipette. Store tubes at -80 °C for further processing.

2. Isolation of total lipids from yeast

NOTE: The following protocol for lipid isolation is based on a well-established and frequently used method that efficiently extracts most neutral lipid species^17,18.
CAUTION: When using organic solvent, always wear appropriate PPE and work inside of a fume hood when possible. During lipid extraction, avoid using plastics that are incompatible with organic solvents. Polypropylene tubes are suitable for the following protocol.

1. Weigh 0.3 g of acid-washed glass beads for each sample and store them in 2 mL microcentrifuge tubes on ice. Remove the cell pellets from the -80 °C freezer and keep them on ice. Add 350 μL methanol and 700 μL chloroform to each sample, resuspend, and transfer to microcentrifuge tubes containing pre-weighed glass beads.
2. Lyse cells by agitating tubes on a vortex 3x for 1 min, with 30 s incubations on ice between agitations. Alternatively, cells can be lysed using a mini bead-beater for three 1-min cycles. Save 25-30 μL of whole cell lysate in a separate tube for scintillation counting.
   NOTE: The saved lysate will be used to determine the relative amount of radioisotope taken up by each sample during the pulse period, which will influence the amount of each sample loaded onto the TLC plate. This is discussed further in step 3.2.
3. Pour the entire contents of the 2 mL microcentrifuge tube into a 15 mL glass centrifuge tube [Tube A]. Wash the 2 mL microcentrifuge tubes by adding 1 mL of methanol and vortexing for 10-15 s. Transfer the 1 mL methanol wash to tube A and add 2 mL of chloroform to tube A followed by 400 μL of water for a final sample volume of 4.45 mL.
4. Vortex samples for 1 min followed by a 5 min centrifugation at 1,000 x g. After centrifugation, the aqueous (upper) and organic (lower) phases should be clearly visually separated with cell debris lying at the interface.
5. Using a glass Pasteur pipette, collect the organic phase from tube A and move to a new 15 mL glass centrifuge tube (Tube B). Add 1 mL of 1M KCl to tube B. To tube A, add 1 mL of methanol and 2 mL of chloroform, and 200 μL MiliQ water. Repeat the vortexing and centrifugation steps on tube A.
6. Once again collect the organic phase from tube A and add it to tube B. Dispose of tube A in an appropriate container. Vortex tube B for 1 min followed by a 5 min centrifugation at 1,000 x g.
7. Remove the upper aqueous layer from tube B and dispose. Add 1 mL of fresh 1 M KCl back to tube B and repeat the vortexing/centrifugation step. Once layers are separated, carefully collect the entire bottom organic layer into a labeled 4 mL glass vial.
   NOTE: At this step, lipid extracts can be stored at -80 °C, or the protocol can be continued for TLC separation of lipids.

3. Separation and quantification of radioisotope-labeled NLs by thin layer chromatography

1. If lipid extracts were placed at -80 °C, slowly bring to room temperature by incubating on ice and subsequently on a benchtop. Completely evaporate solvent from lipid extracts by vacuum drying or using a gentle stream of inert gas (e.g., argon or nitrogen). Meanwhile, pre-heat an oven to 145 °C for heating the TLC plate.
2. Before samples can be loaded onto the TLC plate, determine relative amounts of radiolabel taken up by the cells. Pipette 10 μL of the whole cell lysate from step 2.2 into a 6 mL glass scintillation vial, add 6 mL of scintillation fluid and place vials in a rack. Use a scintillation counter to measure the cpm or dpm of each sample using the count single rack option set to a 1 min counting time. Measure each whole cell lysate in duplicate to obtain an average for each sample. Adjust the loading amount according to a wild type or reference sample
   NOTE: The amount of each sample to load onto the TLC plate can be determined using the following equation: (average sample counts)/(average reference counts) x desired loading volume. For example, if 20 μL of the reference sample is to be loaded onto the TLC plate, and has an average count of 1,000, then an experimental sample with an average count of 2,000 will have 10 μL loaded onto the TLC plate.
3. Reconstitute the sample lipids in 40-50 μL of 1:1 (v/v%) chloroform:methanol by vortexing for 5 min. Prepare 101 mL of the mobile phase solvent in a glass graduated cylinder (see the Representative Results section for an example of major NL species separation by a 50:40:10:1 (v/v/v/v%) Hexane:Petroleum ether:Diethyl ether:Acetic acid solvent).
4. Pour the solvent into a glass TLC chamber containing a 20 x 20 TLC saturation pad and a tight-fitting lid. Prepare a channeled 20 x 20 silica gel 60 G plate by gently marking a line 1.5 cm above the bottom of the plate using a pencil. The line designates the origin and where the lipids will be loaded. Below the line, gently label the sample that will be loaded in each lane. Once the TLC plate has been prepped, incubate the plate in a 145 °C oven for at least 30 min to pre-heat the plate and remove any excess moisture.
5. Once the plate has been sufficiently heated, and the TLC saturation pad is saturated with solvent, remove the TLC plate from the oven and immediately proceed to loading the TLC plate. Loading the plate while it is warm ensures rapid solvent evaporation. For each lipid species of interest, load 5-20 μg of a purified lipid standard onto a lane of the TLC plate to track separation and expected migration distance. Using a pipette, spot 5μL of sample onto the origin of each lane located 1.5 cm above the bottom the of TLC plate. Repeat loading of 5 μL spots until 20-40 μL of sample has been loaded into each lane.
   NOTE: 5-20 μg of unlabeled purified lipids can be added to each sample lane as tracers that can be stained and visualized following TLC separation of lipids. The presence of a stained standard allows for easy tracking and excision of radiolabeled lipid bands for subsequent scintillation counting. Which purified standards are loaded onto the plate will be determined by the NL species of interest. See the Representative Result section for examples of separating oleic acid (FFA), 1,2 dioleoyl-glycerol (DG), triolein (TG), cholesterol (Chol), cholesteryl-linoleate (SE), and squalene in lanes adjacent to the sample lanes.
6. Once the standard and the experimental samples have been loaded, place the plate in the developing chamber and wait until the solvent has reached the top of the plate (40-60 min). Once the plate is fully developed, remove it from the chamber and allow it to dry in the fume hood for 20 minutes.
7. After the plate is dried, cover it with plastic film and place it in a developing cassette with an autoradiography screen. Allow the plate to develop with the screen for 24-48 h.

4. Visualization and quantification of TLC separated lipids

1. Remove the screen from the developing cassette and place inside of a phosphor imager. Select the Phosphor Imaging option and develop at 800-1000 V.
   NOTE: Phosphor imaging gives a qualitative view of radiolabeled lipids on the TLC plate. However, quantification of radiolabeled lipids is best accomplished by scintillation counting, which is described subsequently.
2. Mix 100 mL of p-anisaldehyde reagent (see Supplementary File) and deposit in a glass spray bottle. Spray the TLC plate with p-anisaldehyde reagent until the silica is saturated. Bake the plate in a 145 °C oven for 5 min, or until bands have appeared.
3. To quantify individual lipid species using radiolabel scintillation counting, use a razor blade to scrape the silica gel from the glass TLC plate. Transfer each silica gel band corresponding to a single radiolabeled lipid species to a glass scintillation vial and add 6 mL scintillation fluid. Vortex vigorously until the silica band has been reduced to small pieces.
   1. Alternatively, lipids can be extracted from the silica gel band using the lipid extraction protocol in section 3. If lipids are extracted from the silica gel, evaporate the solvent entirely as in step 3.1, and add 6 mL scintillation fluid to the dried lipids. Place the rack containing scintillation vials into a scintillation counter. Select the Count single rack option and adjust the counting time to 2 min per vial. Results from the scintillation counter will be printed and can be visualized as a bar graph.