The authors would like to thank the members of the Henne lab for help and conceptual advice in the completion of this study. W.M.H. is supported by funds from the Welch Foundation (I-1873), the NIH NIGMS (GM119768), the Ara Paresghian Medical Research Fund, and the UT Southwestern Endowed Scholars Program. S.R has been supported by a T32 program grant (5T32GM008297).

1. Growth and labeling of yeast cells with 14C-acetic acid
<ol>
	<li>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 &#730;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.</li>
	<li>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).</li>
	<li>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 &#176;C for future use.</li>
	<li>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 &#956;Ci/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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Resuspend the cells once more in 500 &#956;L of dextrose-free SC media. Pre-cool a centrifuge equipped for 50 mL conical tubes to -10 &#176;C or on the lowest temperature setting.</li>
	<li>Begin the radiolabeling period by quickly adding 500 &#956;L of radiolabeling media to each 500 &#956;L of cell suspension (final 14C-acetic acid concentration = 5 &#956;Ci/mL). Incubate the tubes in a rotating incubator at 30 &#176;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 &#176;C freezer to a bucket of ice</li>
	<li>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.</li>
	<li>Collect the cell pellet by spinning in a centrifuge at 5,000 x g for 3 min set to -10 &#176;C or on the lowest temperature setting. While sample tubes are spinning, transfer another set of quenching buffer aliquot from the -80 &#176;C freezer to a bucket filled with ice (i.e., one 20 mL tube of quenching buffer per sample).</li>
	<li>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 &#176;C to collect the cells.</li>
	<li>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 &#176;C for further processing.</li>
</ol>
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 species17,18. 
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.
<ol>
	<li>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 &#176;C freezer and keep them on ice. Add 350 &#956;L methanol and 700 &#956;L chloroform to each sample, resuspend, and transfer to microcentrifuge tubes containing pre-weighed glass beads.</li>
	<li>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 &#956;L 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.</li>
	<li>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 &#956;L of water for a final sample volume of 4.45 mL.</li>
	<li>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.</li>
	<li>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 &#956;L MiliQ water. Repeat the vortexing and centrifugation steps on tube A.</li>
	<li>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.</li>
	<li>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 &#176;C, or the protocol can be continued for TLC separation of lipids.</li>
</ol>
3. Separation and quantification of radioisotope-labeled NLs by thin layer chromatography
<ol>
	<li>If lipid extracts were placed at -80 &#176;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 &#176;C for heating the TLC plate.</li>
	<li>Before samples can be loaded onto the TLC plate, determine relative amounts of radiolabel taken up by the cells. Pipette 10 &#956;L 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 &#956;L 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 &#956;L loaded onto the TLC plate.</li>
	<li>Reconstitute the sample lipids in 40-50 &#956;L 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).</li>
	<li>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 &#176;C oven for at least 30 min to pre-heat the plate and remove any excess moisture.</li>
	<li>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 &#956;g of a purified lipid standard onto a lane of the TLC plate to track separation and expected migration distance. Using a pipette, spot 5&#956;L of sample onto the origin of each lane located 1.5 cm above the bottom the of TLC plate. Repeat loading of 5 &#956;L spots until 20-40 &#956;L of sample has been loaded into each lane. 
	NOTE: 5-20 &#956;g 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&#160;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.</li>
	<li>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.</li>
	<li>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.</li>
</ol>
4. Visualization and quantification of TLC separated lipids
<ol>
	<li>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.</li>
	<li>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 &#176;C oven for 5 min, or until bands have appeared.</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
</ol>

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The authors declare there are no competing interests in the preparation of this manuscript.

Here, a versatile radiolabeling protocol to quantitatively monitor the synthesis of NL species in yeast is presented. This protocol is very modular, which allows for the procedure to be finished within 3-6 days. Additionally, a wealth of literature exists on the use of TLC to separate lipid species and metabolites, which should permit the user to detect several lipid species of interest with a simple change of TLC solvent systems16,19.This protocol is conducive t...

Acetyl-CoA is the fundamental building block of diverse biomolecules including neutral lipids (NLs), which serve as a versatile biomolecular currency for building membranes, generating ATP, and regulating cell signaling1,2. The availability of NLs to be shunted into any of these respective pathways is, in part, regulated by their storage. Lipid droplets (LDs), cytoplasmic organelles composed of hydrophobic cores of triglycerides (TGs) and sterol-esters (SEs), are the main storage compartments of most cellular NLs. As such, LDs sequester and regulate NLs, which can be degraded and subsequently utilized for biochemical and metabolic processes3,4. It is known that the mis-regulation of NL and LD-associated proteins is correlated with the onset of pathologies including lipodystrophy and metabolic syndromes5,6. Because of this, current LD research is intensely focused on how NL synthesis is regulated spatially, temporally, and across distinct tissues of multi-cellular organisms. Due to the ubiquitous cellular roles for NLs, many enzymes responsible for the synthesis and regulation of NLs are conserved throughout eukaryotes7. Indeed, even some prokaryotes store NLs in LDs8. Therefore, genetically tractable model organisms such as Saccharomyces cerevisiae (budding yeast) have been useful for the study of NL synthesis and regulation.
The separation and quantification of NLs from cell extracts can be accomplished in a myriad of ways, including gas chromatography-mass spectrometry (GC-MS), high-performance liquid chromatography (HPLC), and ultra-performance liquid chromatography-mass spectrometry (UPLC-MS)9,10,11. Perhaps the simplest method for separating NLs is via thin layer chromatography (TLC), which allows for subsequent densitometric quantification from a standard curve12,13. Although TLC provides only a course-grained separation of NLs, it remains a powerful technique because it is inexpensive, and it allows for the rapid separation of NLs from several samples simultaneously. Two of the most considerable challenges facing the study of NLs via TLC are: 1) the broad range of cellular abundances of NL species and their intermediates, and 2) the range of hydrophilicity/hydrophobicity of lipid intermediates within NL synthesis pathways. Consequently, the quantification of NL species via TLC is typically restricted to the most abundant species; however, introduction of a 14C-acetic acid radiolabel can significantly enhance the detection of low abundance intermediates within NL pathways. Acetic acid is rapidly converted into acetyl-CoA by the acetyl-CoA synthetase ACS214, which makes 14C-acetic acid a suitable radiolabeling substrate in yeast15. Additionally, separation of both hydrophobic NLs and hydrophilic intermediates of NLs can be achieved by TLC through the use of multiple solvent systems16. Here, a method is presented for the separation of NLs using 14C-acetic acid metabolic labeling in yeast. Lipids labeled during the pulse period are subsequently isolated by a well-established total lipid isolation protocol17, followed by the separation of NL species by TLC. Developing of TLC plates by both autoradiography to visualize labeled lipids, and a chemical spray to visualize total lipids, permits for multiple methods of quantification. Individual lipid bands can also be easily extracted from the TLC plate using a razor blade, and scintillation counting can be used to quantify amount of radiolabeled material within the band.

<table><tbody><tr><td>[1-C14] Acetic acid sodium salt specific activity: 45-60mCi</td><td>PerkinElmer</td><td>NEC084H001MC</td><td></td></tr><tr><td>18:1 1,2 dioleoyl-sn-glycerol</td><td>Avanti</td><td>800811O</td><td></td></tr><tr><td>200 proof absolute ethanol</td><td>Sigma</td><td>459836</td><td></td></tr><tr><td>Acid washed glass beads 425-600um</td><td>Sigma</td><td>G8772</td><td></td></tr><tr><td>Amber bulbs for Pastuer pipettes</td><td>Fisher</td><td>03-448-24</td><td></td></tr><tr><td>Ammonium Sulfate &gt;99%</td><td>Sigma</td><td>A4418</td><td></td></tr><tr><td>Beckman LS6500 scintillation counter</td><td>PerkinElmer</td><td>A481000</td><td></td></tr><tr><td>Chloroform (HPLC grade)</td><td>Fisher</td><td>C607SK</td><td></td></tr><tr><td>Cholesterol &gt;99%</td><td>Sigma</td><td>C8667</td><td></td></tr><tr><td>Cholesteryl-linoleate &gt;98%</td><td>Sigma</td><td>C0289</td><td></td></tr><tr><td>Concentrated sulfuric acid</td><td>Sigma</td><td>339741</td><td></td></tr><tr><td>Corning 50mL conical tubes, polypropylene with centristar cap</td><td>Sigma</td><td>CLS430829</td><td></td></tr><tr><td>Dextrose, anhydrous grade</td><td>Sigma</td><td>D9434</td><td></td></tr><tr><td>Diethyl ether anhydrous grade</td><td>Sigma</td><td>296082</td><td></td></tr><tr><td>Drying oven</td><td>Fisher</td><td>11-475-155</td><td></td></tr><tr><td>EcoLume scintillation liquid</td><td>VWR</td><td>IC88247001</td><td></td></tr><tr><td>Eppendorf 5424R centrifuge</td><td>Fisher</td><td>05-401-205</td><td></td></tr><tr><td>GE Storage phosphor screen</td><td>Sigma</td><td>GE28-9564-75</td><td></td></tr><tr><td>GE Typhoon FLA9500 imager</td><td></td><td></td><td></td></tr><tr><td>Glacial acetic acid, ACS grade</td><td>Sigma</td><td>695092</td><td></td></tr><tr><td>Glass 6mL scintillation vials</td><td>Sigma</td><td>M1901</td><td></td></tr><tr><td>Glass centrifuge tube caps</td><td>Fisher</td><td>14-595-36A</td><td></td></tr><tr><td>Glass centrifuge tubes</td><td>Fisher</td><td>14-595-35A</td><td></td></tr><tr><td>Glass Pasteur pipette</td><td>Fisher</td><td>13-678-20C</td><td></td></tr><tr><td>Hexane, anhydrous grade</td><td>Sigma</td><td>296090</td><td></td></tr><tr><td>L-Adenine &gt;99%</td><td>Sigma</td><td>A8626</td><td></td></tr><tr><td>L-Alanine &gt;98%</td><td>Sigma</td><td>A7627</td><td></td></tr><tr><td>L-Arginine &gt;99%</td><td>Sigma</td><td>A1270000</td><td></td></tr><tr><td>L-Asparagine &gt;98%</td><td>Sigma</td><td>A0884</td><td></td></tr><tr><td>L-Aspartate &gt;98%</td><td>Sigma</td><td>A9256</td><td></td></tr><tr><td>L-Cysteine &gt;97%</td><td>Sigma</td><td>W326305</td><td></td></tr><tr><td>L-Glutamic acid monosodium salt monohydrate &gt;98%</td><td>Sigma</td><td>49621</td><td></td></tr><tr><td>L-Glutamine &gt;99%</td><td>Sigma</td><td>G3126</td><td></td></tr><tr><td>L-Glycine &gt;99%</td><td>Sigma</td><td>G8898</td><td></td></tr><tr><td>L-Histidine &gt;99%</td><td>Sigma</td><td>H8000</td><td></td></tr><tr><td>L-Isoleucine &gt;98%</td><td>Sigma</td><td>I2752</td><td></td></tr><tr><td>L-Leucine &gt;98%</td><td>Sigma</td><td>L8000</td><td></td></tr><tr><td>L-Lysine &gt;98%</td><td>Sigma</td><td>L5501</td><td></td></tr><tr><td>L-Methionine, HPLC grade</td><td>Sigma</td><td>M9625</td><td></td></tr><tr><td>L-Phenylalanine, reagent grade</td><td>Sigma</td><td>P2126</td><td></td></tr><tr><td>L-Proline &gt;99%</td><td>Sigma</td><td>P0380</td><td></td></tr><tr><td>L-Serine &gt;99%</td><td>Sigma</td><td>S4500</td><td></td></tr><tr><td>L-Theronine, reagent grade</td><td>Sigma</td><td>T8625</td><td></td></tr><tr><td>L-Tryptophan &gt;98%</td><td>Sigma</td><td>T0254</td><td></td></tr><tr><td>L-Tyrosine &gt;98%</td><td>Sigma</td><td>T3754</td><td></td></tr><tr><td>L-Uracil &gt;99%</td><td>Sigma</td><td>U0750</td><td></td></tr><tr><td>L-Valine &gt;98%</td><td>Sigma</td><td>V0500</td><td></td></tr><tr><td>Methanol, ACS grade</td><td>Fisher</td><td>A412</td><td></td></tr><tr><td>Oleic acid &gt;99%</td><td>Sigma</td><td>O1008</td><td></td></tr><tr><td>p-anisaldehyde</td><td>Sigma</td><td>A88107</td><td></td></tr><tr><td>Petroleum ether, ACS grade</td><td>Sigma</td><td>184519</td><td></td></tr><tr><td>Phosphatidylcholine, dipalmitoyl &gt;99%</td><td>Sigma</td><td>P1652</td><td></td></tr><tr><td>Pipettes</td><td>Eppendorf</td><td>2231000713</td><td></td></tr><tr><td>Potassium chloride, ACS grade</td><td>Sigma</td><td>P3911</td><td></td></tr><tr><td>Sodium Hydroxide pellets, certified ACS</td><td>Fisher</td><td>S318-100</td><td></td></tr><tr><td>Squalene &gt;98%</td><td>Sigma</td><td>S3626</td><td></td></tr><tr><td>Succinic Acid crystalline/certified</td><td>Fisher</td><td>110-15-6</td><td></td></tr><tr><td>TLC saturation pad</td><td>Sigma</td><td>Z265225</td><td></td></tr><tr><td>TLC silica gel 60G glass channeled plate</td><td>Fisher</td><td>NC9825743</td><td>No fluorescent indicators</td></tr><tr><td>Transparency plastic film</td><td>Apollo</td><td>829903</td><td></td></tr><tr><td>Tricine</td><td>Sigma</td><td>T0377</td><td></td></tr><tr><td>Triolein &gt;99%</td><td>Sigma</td><td>T7140</td><td></td></tr><tr><td>Vortex mixer</td><td>Fisher</td><td>02-215-414</td><td></td></tr><tr><td>Whatman exposure cassette</td><td>Sigma</td><td>WHA29175523</td><td></td></tr><tr><td>Yeast nitrogen base without ammonium sulfate and amino acids</td><td>Sigma</td><td>Y1251</td><td></td></tr></tbody></table>

analysis of neutral lipid synthesis in saccharomyces cerevisiae by metabolic labeling and thin layer chromatography

Neutral lipids (NLs) are a class of hydrophobic, chargeless biomolecules that play key roles in energy and lipid homeostasis. NLs are synthesized de novo&#160;from acetyl-CoA and are primarily present in eukaryotes in the form of triglycerides (TGs) and sterol-esters (SEs). The enzymes responsible for the synthesis of NLs are highly conserved from Saccharomyces cerevisiae (yeast) to humans, making yeast a useful model organism to dissect the function and regulation of NL metabolism enzymes. While much is known about how acetyl-CoA is converted into a diverse set of NL species, mechanisms for regulating NL metabolism enzymes, and how mis-regulation can contribute to cellular pathologies, are still being discovered. Numerous methods for the isolation and characterization of NL species have been developed and used over decades of research; however, a quantitative and simple protocol for the comprehensive characterization of major NL species has not been discussed. Here, a simple and adaptable method to quantify the de novo&#160;synthesis of major NL species in yeast is presented. We apply 14C-acetic acid metabolic labeling coupled with thin layer chromatography to separate and quantify a diverse range of physiologically important NLs. Additionally, this method can be easily applied to study in vivo reaction rates of NL enzymes or degradation of NL species over time.

In this protocol, we have demonstrated that the labeling, detection, and quantification of NL species can be accomplished by 14C-acetic acid metabolic labeling. Major NL species can be separated in a solvent system of 50:40:10:1 (v/v/v/v%) Hexane:Petroleum ether:Diethyl ether:Acetic acid (Figure 1A,B). Phosphor imaging allows for visualization of labeled free fatty acid (FFA), triacylglycerol (TG), diacylglycerol (DG), cholesterol (Chol), and squalene (SQ) (<stron...

Watch this Scientific Journal Video about Analysis of Neutral Lipid Synthesis in Saccharomyces cerevisiae by Metabolic Labeling and Thin Layer Chromatography at JoVE.com

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

Here, a protocol is presented for the metabolic labeling of yeast with 14C-acetic acid, which is coupled with thin layer chromatography for the separation of neutral lipids.

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

Neutral lipids (NLs) are a class of hydrophobic, chargeless biomolecules that play key roles in energy and lipid homeostasis. NLs are synthesized de ...

analysis-neutral-lipid-synthesis-saccharomyces-cerevisiae-metabolic

Research

JoVE Journal

Biochemistry

3.0K Views.  University of Texas Southwestern Medical Center. Deze techniek maakt gebruik van instrumenten die vaak in de meeste laboratoriumomgevingen worden aangetroffen en maakt het mogelijk om meerdere monsters tegelijkertijd te analyseren. Deze methode kan belangrijke inzichten bieden in hoe lipidenmetabolisme enzymen worden gereguleerd in een cellulaire context en in verschillende genetische achtergronden. Ent een kolonie van een gistkweekplaat in 20 milliliter SC-media met 2% dextrose en incubeer bij 30 graden Celsius voor een nacht met schudden ...

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

Lipid Index Determination by Liquid Fluorescence Recovery in the Fungal Pathogen Ustilago Maydis

The article shows how to implement the LD index assay, which is a sensitive microplate assay to determine the accumulation of triacylglycerols (TAGs) in lipid droplets (LDs). LD index is obtained without lipid extraction. It allows measuring the LDs content in high-throughput experiments under different conditions such as growth in rich or nitrogen depleted media. Albeit the method was described for the first time to study the lipid droplet metabolism in Saccharomyces cerevisiae, it was successfully applied to the basidiomycete Ustilago maydis. Interestingly, and because LDs are organelles phylogenetically conserved in eukaryotic cells, the method can be applied to a large variety of cells, from yeast to mammalian cells. The LD index is based on the liquid fluorescence recovery assay (LFR) of the BODIPY 493/503 under quenching conditions, by the addition of cells fixed with formaldehyde. Potassium iodine is used as a fluorescence quencher. The ratio between the fluorescence and the optical density slopes is named LD index. Slopes are calculated from the straight lines obtained when BODIPY fluorescence and optical density at 600 nm (OD600) are plotted against sample addition. Optimal data quality is reflected by correlation coefficients equal or above 0.9 (r &#8805; 0.9). Multiple samples can be read simultaneously as it can be implemented in a microplate. Since BODIPY 493/503 is a lipophilic fluorescent dye that partitions into the lipid droplets, it can be used in many types of cells that accumulate LDs.

Lipid Index Determination by Liquid Fluorescence Recovery in the Fungal Pathogen Ustilago Maydis

Here, we describe a protocol to obtain the lipid droplet index (LD index) to study the dynamics of triacylglycerols in cells cultured in high-throughput experiments. The LD index assay is an easy and reliable method that uses BODIPY 493/503. This assay does not need dispendious lipid extraction or microscopy analysis.

The article shows how to implement the LD index assay, which is a sensitive microplate assay to determine the accumulation of triacylglycerols (TAGs) in ...

Saccharomyces cerevisiae Metabolic Labeling with 4-thiouracil and the Quantification of Newly Synthesized mRNA As a Proxy for RNA Polymerase II Activity

Global defects in RNA polymerase II transcription might be overlooked by transcriptomic studies analyzing steady-state RNA. Indeed, the global decrease in mRNA synthesis has been shown to be compensated by a simultaneous decrease in mRNA degradation to restore normal steady-state levels. Hence, the genome-wide quantification of mRNA synthesis, independently from mRNA decay, is the best direct reflection of RNA polymerase II transcriptional activity. Here, we discuss a method using non-perturbing metabolic labeling of nascent RNAs in Saccharomyces cerevisiae (S. cerevisiae). Specifically, the cells are cultured for 6 min with a uracil analog, 4-thiouracil, and the labeled newly transcribed RNAs are purified and quantified to determine the synthesis rates of all individual mRNA. Moreover, using labeled Schizosaccharomyces pombe cells as internal standard allows comparing mRNA synthesis in different S. cerevisiae strains. Using this protocol and fitting the data with a dynamic kinetic model, the corresponding mRNA decay rates can be determined.

Saccharomyces cerevisiae Metabolic Labeling with 4-thiouracil and the Quantification of Newly Synthesized mRNA As a Proxy for RNA Polymerase II Activity

The protocol described here is based on the genome-wide quantification of newly synthesized mRNA purified from yeast cells labeled with 4-thiouracil. This method allows to measure mRNA synthesis uncoupled from mRNA decay and, thus, provides an accurate measurement of RNA polymerase II transcription.

Global defects in RNA polymerase II transcription might be overlooked by transcriptomic studies analyzing steady-state RNA. Indeed, the global decrease in ...

Genetics

Mass Spectrometric Analysis of Glycosphingolipid Antigens

Glycosphingolipids (GSL's) belong to the glycoconjugate class of biomacromolecules, which bear structural information for significant biological processes such as embryonic development, signal transduction, and immune receptor recognition1-2. They contain complex sugar moieties in the form of isomers, and lipid moieties with variations including fatty acyl chain length, unsaturation, and hydroxylation. Both carbohydrate and ceramide portions may be basis of biological significance. For example, tri-hexosylceramides include globotriaosylceramide (Gal&alpha;4Gal&beta;4Glc&beta;1Cer) and isoglobotriaosylceramide (Gal&alpha;3Gal&beta;4Glc&beta;1Cer), which have identical molecular masses but distinct sugar linkages of carbohydrate moiety, responsible for completely different biological functions3-4. In another example, it has been demonstrated that modification of the ceramide part of alpha-galactosylceramide, a potent agonist ligand for invariant NKT cells, changes their cytokine secretion profiles and function in animal models of cancer and auto-immune diseases5. The difficulty in performing a structural analysis of isomers in immune organs and cells serve as a barrier for determining many biological functions6.
Here, we present a visualized version of a method for relatively simple, rapid, and sensitive analysis of glycosphingolipid profiles in immune cells7-9. This method is based on extraction and chemical modification (permethylation, see below Figure 5A, all OH groups of hexose were replaced by MeO after permethylation reaction) of glycosphingolipids10-15, followed by subsequent analysis using matrix-assisted laser desorption/ionization time-of-flight mass spectrometry (MALDI-TOF/MS) and ion trap mass spectrometry. This method requires 50 million immune cells for a complete analysis. The experiments can be completed within a week. The relative abundance of the various glycosphingolipids can be delineated by comparison to synthetic standards. This method has a sensitivity of measuring 1% iGb3 among Gb3 isomers, when 2 fmol of total iGb3/Gb3 mixture is present9.
Ion trap mass spectrometry can be used to analyze isomers. For example, to analyze the presence of globotriaosylceramide and isoglobtriaosylceramide in the same sample, one can use the fragmentation of glycosphingolipid molecules to structurally discriminate between the two (see below Figure 5). Furthermore, chemical modification of the sugar moieties (through a permethylation reaction) improves the ionization and fragmentation efficiencies for higher sensitivity and specificity, and increases the stability of sialic acid residues. The extraction and chemical modification of glycosphingolipids can be performed in a classic certified chemical hood, and the mass spectrometry can be performed by core facilities with ion trap MS instruments.

A specific and sensitive method to gain insight into the expression profile of glycosphingolipid antigens in immune organs and cells is described. The method takes advantage of the ion trap mass spectrometry allowing step-wise fragmentation of glycosphingolipid molecules for structural analysis in comparison to chemically synthesized standards.

Glycosphingolipids (GSL's) belong to the glycoconjugate class of biomacromolecules, which bear structural information for significant biological processes ...

Immunology and Infection

Identification of protein complexes with quantitative proteomics in S. cerevisiae

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as important intracellular signalling molecules. However, unlike proteins, lipids are small hydrophobic molecules that traffic primarily by poorly described nonvesicular routes, which are hypothesized to occur at membrane contact sites (MCSs). MCSs are regions where the endoplasmic reticulum (ER) makes direct physical contact with a partnering organelle, e.g., plasma membrane (PM). The ER portion of ER-PM MCSs is enriched in lipid-synthesizing enzymes, suggesting that lipid synthesis is directed to these sites and implying that MCSs are important for lipid traffic. Yeast is an ideal model to study ER-PM MCSs because of their abundance, with over 1000 contacts per cell, and their conserved nature in all eukaryotes. Uncovering the proteins that constitute MCSs is critical to understanding how lipids traffic is accomplished in cells, and how they act as signaling molecules. We have found that an ER called Scs2p localize to ER-PM MCSs and is important for their formation. We are focused on uncovering the molecular partners of Scs2p. Identification of protein complexes traditionally relies on first resolving purified protein samples by gel electrophoresis, followed by in-gel digestion of protein bands and analysis of peptides by mass spectrometry. This often limits the study to a small subset of proteins. Also, protein complexes are exposed to denaturing or non-physiological conditions during the procedure. To circumvent these problems, we have implemented a large-scale quantitative proteomics technique to extract unbiased and quantified data. We use stable isotope labeling with amino acids in cell culture (SILAC) to incorporate staple isotope nuclei in proteins in an untagged control strain. Equal volumes of tagged culture and untagged, SILAC-labeled culture are mixed together and lysed by grinding in liquid nitrogen. We then carry out an affinity purification procedure to pull down protein complexes. Finally, we precipitate the protein sample, which is ready for analysis by high-performance liquid chromatography/ tandem mass spectrometry. Most importantly, proteins in the control strain are labeled by the heavy isotope and will produce a mass/ charge shift that can be quantified against the unlabeled proteins in the bait strain. Therefore, contaminants, or unspecific binding can be easily eliminated. By using this approach, we have identified several novel proteins that localize to ER-PM MCSs. Here we present a detailed description of our approach. 

Here we describe a new quantitative proteomics technique for identifying protein complexes in Saccharomyces cerevisiae. In this study, we have used the SILAC method together with affinity purification followed by tandem mass spectrometry to identify with high specificity the binding partners of an ER protein, Scs2p.

Lipids are the building blocks of cellular membranes that function as barriers and in compartmentalization of cellular processes, and recently, as ...

Biology

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of biological membranes, energy storage and production, cellular signaling, vesicular transport of proteins, organelle biogenesis, and regulated cell death. Because the budding yeast Saccharomyces cerevisiae is a unicellular eukaryote amenable to thorough molecular analyses, its use as a model organism helped uncover mechanisms linking lipid metabolism and intracellular transport to complex biological processes within eukaryotic cells. The availability of a versatile analytical method for the robust, sensitive, and accurate quantitative assessment of major classes of lipids within a yeast cell is crucial for getting deep insights into these mechanisms. Here we present a protocol to use liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of major cellular lipids of S. cerevisiae. The LC-MS/MS method described is versatile and robust. It enables the identification and quantification of numerous species (including different isobaric or isomeric forms) within each of the 10 lipid classes. This method is sensitive and allows identification and quantitation of some lipid species at concentrations as low as 0.2 pmol/&#181;L. The method has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. The use of alternative mobile phase additives for electrospray ionization mass spectrometry in this method can increase the efficiency of ionization for some lipid species and can be therefore used to improve their identification and quantitation.

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol using liquid chromatography coupled with tandem mass spectrometry to identify and quantify major cellular lipids in Saccharomyces cerevisiae. The described method for a quantitative assessment of major lipid classes within a yeast cell is versatile, robust, and sensitive.

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of ...

Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

This protocol describes the growth and stimulation, with the fatty acid oleate, of isotopically heavy and light S. cerevisiae cells. Cells are ground using a cryolysis procedure in a ball mill grinder and the resulting grindate brought into solution by urea solubilization. This procedure allows for the lysis of the cells in a metabolically inactive state, preserving phosphorylation and preventing reorientation of the phosphoproteome during cell lysis.  Following reduction, alkylation, trypsin digestion of the proteins, the samples are desalted on C18 columns and the sample complexity reduced by fractionation using hydrophilic interaction chromatography (HILIC). HILIC columns preferentially retain hydrophilic molecules which is well suited for phosphoproteomics.  Phosphorylated peptides tend to elute later in the chromatographic profile than the non phosphorylated counterparts. After fractionation, phosphopeptides are enriched using immobilized metal chromatography, which relies on charge-based affinities for phosphopeptide enrichment. At the end of this procedure the samples are ready to be quantitatively analyzed by mass spectrometry.

Quantitative Phosphoproteomics in Fatty Acid Stimulated Saccharomyces cerevisiae

Description of a quantitative phosphorylation procedure using cryolysis, urea solubilziation, HILIC fractionation and IMAC enrichment of phosphorylated peptides.

This protocol describes the growth and stimulation, with the fatty acid oleate, of isotopically heavy and light S. cerevisiae cells. Cells are ground ...

A Quantitative Assessment of The Yeast Lipidome using Electrospray Ionization Mass Spectrometry

Lipids are one of the major classes of biomolecules and play important roles membrane dynamics, energy storage, and signalling1-4. The budding yeast Saccharomyces cerevisiae, a genetically and biochemically manipulable unicellular eukaryote with annotated genome and very simple lipidome, is a valuable model for studying biological functions of various lipid species in multicellular eukaryotes2,3,5. S. cerevisiae has 10 major classes of lipids with chain lengths mainly of 16 or 18 carbon atoms and either zero or one degree of unsaturation6,7. Existing methods for lipid identification and quantification - such as high performance liquid chromatography, thin-layer chromatography, fluorescence microscopy, and gas chromatography followed by MS - are well established but have low sensitivity, insufficiently separate various molecular forms of lipids, require lipid derivitization prior to analysis, or can be quite time consuming. Here we present a detailed description of our experimental approach to solve these inherent limitations by using survey-scan ESI/MS for the identification and quantification of the entire complement of lipids in yeast cells. The described method does not require chromatographic separation of complex lipid mixtures recovered from yeast cells, thereby greatly accelerating the process of data acquisition. This method enables lipid identification and quantification at the concentrations as low as g/ml and has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. Lipids extraction from whole yeast cells for using this method of lipid analysis takes two to three hours. It takes only five to ten minutes to run each sample of extracted and dried lipids on a Q-TOF mass spectrometer equipped with a nano-electrospray source.

We describe a new quantitative lipidomics method for identifying numerous lipid species in yeast using survey-scan electrospray ionization mass spectrometry (ESI/MS). This method exceeds currently available methods for lipid identification and quantification in the ability to resolve various molecular forms of lipids, sensitivity, and speed.

Lipids are one of the major classes of biomolecules and play important roles membrane dynamics, energy storage, and signalling1-4. The budding yeast ...

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

Mitochondria are essential organelles of eukaryotic cells capable of aerobic respiration. They contain circular genome and gene expression apparatus. A mitochondrial genome of baker&#8217;s yeast encodes eight proteins: three subunits of the cytochrome c oxidase (Cox1p, Cox2p, and Cox3p), three subunits of the ATP synthase (Atp6p, Atp8p, and Atp9p), a subunit of the ubiquinol-cytochrome c oxidoreductase enzyme, cytochrome b (Cytb), and mitochondrial ribosomal protein Var1p. The purpose of the method described here is to specifically label these proteins with 35S methionine, separate them by electrophoresis and visualize the signals as discrete bands on the screen. The procedure involves several steps. First, yeast cells are cultured in a galactose-containing medium until they reach the late logarithmic growth stage. Next, cycloheximide treatment blocks cytoplasmic translation and allows 35S methionine incorporation only in mitochondrial translation products. Then, all proteins are extracted from yeast cells and separated by polyacrylamide gel electrophoresis. Finally, the gel is dried and incubated with the storage phosphor screen. The screen is scanned on a phosphorimager revealing the bands. The method can be applied to compare the biosynthesis rate of a single polypeptide in the mitochondria of a mutant yeast strain versus the wild type, which is useful for studying mitochondrial gene expression defects. This protocol gives valuable information about the translation rate of all yeast mitochondrial mRNAs. However, it requires several controls and additional experiments to make proper conclusions.

Labelling and Visualization of Mitochondrial Genome Expression Products in Baker's Yeast Saccharomyces cerevisiae

Baker&#8217;s yeast mitochondrial genome encodes eight polypeptides. The goal of the current protocol is to label all of them and subsequently visualize them as separate bands.

Mitochondria are essential organelles of eukaryotic cells capable of aerobic respiration. They contain circular genome and gene expression apparatus. A ...

Quantitative Metabolomics of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Metabolomics is a methodology used for the identification and quantification of many low-molecular-weight intermediates and products of metabolism within a cell, tissue, organ, biological fluid, or organism. Metabolomics traditionally focuses on water-soluble metabolites. The water-soluble metabolome is the final product of a complex cellular network that integrates various genomic, epigenomic, transcriptomic, proteomic, and environmental factors. Hence, the metabolomic analysis directly assesses the outcome of the action for all these factors in a plethora of biological processes within various organisms. One of these organisms is the budding yeast Saccharomyces cerevisiae, a unicellular eukaryote with the fully sequenced genome. Because S. cerevisiae is amenable to comprehensive molecular analyses, it is used as a model for dissecting mechanisms underlying many biological processes within the eukaryotic cell. A versatile analytical method for the robust, sensitive, and accurate quantitative assessment of the water-soluble metabolome would provide the essential methodology for dissecting these mechanisms. Here we present a protocol for the optimized conditions of metabolic activity quenching in and water-soluble metabolite extraction from S. cerevisiae cells. The protocol also describes the use of liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of the extracted water-soluble metabolites. The LC-MS/MS method of non-targeted metabolomics described here is versatile and robust. It enables the identification and quantification of more than 370 water-soluble metabolites with diverse structural, physical, and chemical properties, including different structural isomers and stereoisomeric forms of these metabolites. These metabolites include various energy carrier molecules, nucleotides, amino acids, monosaccharides, intermediates of glycolysis, and tricarboxylic cycle intermediates. The LC-MS/MS method of non-targeted metabolomics is sensitive and allows the identification and quantitation of some water-soluble metabolites at concentrations as low as 0.05 pmol/&#181;L. The method has been successfully used for assessing water-soluble metabolomes of wild-type and mutant yeast cells cultured under different conditions.

Quantitative Metabolomics of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol for the identification and quantitation of major classes of water-soluble metabolites in the yeast Saccharomyces cerevisiae. The described method is versatile, robust, and sensitive. It allows the separation of structural isomers and stereoisomeric forms of water-soluble metabolites from each other.

Metabolomics is a methodology used for the identification and quantification of many low-molecular-weight intermediates and products of metabolism within ...

Workflow Based on the Combination of Isotopic Tracer Experiments to Investigate Microbial Metabolism of Multiple Nutrient Sources

Studies in the field of microbiology rely on the implementation of a wide range of methodologies. In particular, the development of appropriate methods substantially contributes to providing extensive knowledge of the metabolism of microorganisms growing in chemically defined media containing unique nitrogen and carbon sources. In contrast, the management through metabolism of multiple nutrient sources, despite their broad presence in natural or industrial environments, remains virtually unexplored. This situation is mainly due to the lack of suitable methodologies, which hinders investigations.
We report an experimental strategy to quantitatively and comprehensively explore how metabolism operates when a nutrient is provided as a mixture of different molecules, i.e., a complex resource. Here, we describe its application for assessing the partitioning of multiple nitrogen sources through the yeast metabolic network. The workflow combines information obtained during stable isotope tracer experiments using selected 13C- or 15N-labeled substrates. It first consists of parallel and reproducible fermentations in the same medium, which includes a mixture of N-containing molecules; however,a selected nitrogen source is labeled each time. A combination of analytical procedures (HPLC, GC-MS) is implemented to assess the labeling patterns of targeted compounds and to quantify the consumption and recovery of substrates in other metabolites. An integrated analysis of the complete dataset provides an overview of the fate of consumed substrates within cells. This approach requires an accurate protocol for the collection of samples&#8211;facilitated by a robot-assisted system for online monitoring of fermentations&#8211;and the achievement of numerous time-consuming analyses. Despite these constraints, it allowed understanding, for the first time, the partitioning of multiple nitrogen sources throughout the yeast metabolic network. We elucidated the redistribution of nitrogen from more abundant sources toward other N-compounds and determined the metabolic origins of volatile molecules and proteinogenic amino acids.

This protocol describes an experimental procedure to quantitatively and comprehensively investigate the metabolism of multiple nutrient sources. This workflow, based on a combination of isotopic tracer experiments and an analytical procedure, allows the fate of consumed nutrients and the metabolic origin of molecules synthetized by microorganisms to be determined.

Studies in the field of microbiology rely on the implementation of a wide range of methodologies. In particular, the development of appropriate methods ...

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

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