MUSC Department of Biochemistry & Molecular Biology
Website compiled by Stephen Krombach, Ben Pettus and Lewis W. Stillway.
Before TLC can be performed, lipids must first be extracted in an undegraded and uncontaminated state. The most common method of lipid extraction was developed by Bligh and Dyer and is a simplified version of the classical Folch procedure. The extract can then be analyzed and normalized to lipid phosphate.
Extraction of sphingolipids using the method of Bligh and Dyer:
1.) Grow 2-5 x 106 cells per sample to be analyzed. Treat as desired. Adherent cells need not be suspended.
2.) Pellet cells by centrifugation at 1500 rpm for 5 minutes. See Picture
3.) Aspirate off the media or wash PBS (phosphate buffered saline) from the pellet.
4.) Resuspend pellet in 3 ml of chloroform:methanol (1:2 pre-mixed) and agitate (using a Vortex-Genie) until an even suspension is attained. See Picture
5.) Add 0.8 ml of water and agitate. Transfer the resuspended cells to a glass screw-cap tube and let them set on the bench for 30 minutes (or overnight at 4oC for best extraction). If there is a phase break at this point, it can be corrected by the addition of 0.5 ml of methanol. A premature phase break can hinder proper extraction.
6.) Pellet the cellular debris via table top centrifugation at 3000 rpm for 5 minutes. Transfer liquid material to a fresh tube and discard cellular debris.
7.) Add 1 ml of chloroform and 1 ml of water (See Picture) and vortex. These additions will induce a break of liquid material into an organic (lower) and aqueous (upper) phase. Allow 15 to 30 minutes for the phases to separate and then centrifuge for 5 minutes at 3000 rpm to obtain clean phase separation.
8.) Transfer the lower organic phase directly to a new tube.
9.) Evaporate the solvent from the extracted lipids in the 2 ml of chloroform via a speed vacuum apparatus (See Picture) or under a stream of dry nitrogen.
10.) Resuspend the lipids in chloroform with one aliquot designated for phosphate measurement (1/3), another aliquot for experimental measurement (1/3), and a final aliquot as a backup.
Lipids should be stored as either a powder or in chloroform at -20oC.
Lipid Phosphate Measurement
This method is used to more accurately quantitate sphingolipid samples. It normalizes for variances in cell number and extraction efficiency.
1.) Prepare a standard curve of phosphates in duplicate. Put 0,3,5,7,10,12,15,20,30 microliters of 1mM NaH2PO4 (1nM per microliter) in labeled tubes. See Picture
2.) Evaporate the solvent by speed vacuum (See Picture) or under a stream of nitrogen from the aliquots obtained by the extraction above. Use the same type of glass tube for both the standards and the samples to ensure consistent ashing.
3.) Add 0.6 ml of ashing buffer consisting of H2O:10N H2SO4: 70% HClO4 (40:9:1) and agitate (vortex).
4.) Place samples in a heating block at 160oC overnight (See Picture). Approximately 100 microliters should be left in the tube after incubation.
5.) Add 0.9 ml of water and agitate (vortex) after allowing samples to cool.
6.) Add 0.5 ml of ammonium molybdate (0.9% w/v) and add 0.2 ml of freshly prepared L-ascorbic acid (10% w/v). Agitate (vortex) tubes very vigorously.
7.) Incubate at 45oC in a water bath for 30 minutes. After incubation a change in color will be apparent (See Picture).
8.) Read samples on a spectrophotometer at 820 nm (See Picture) in 0.8 ml cuvettes. First construct a standard curve and then use it to calculate experimental phosphate values.
TLC Procedure
1.) The solvent tank to be used must be equilibrated with the solvent used for lipid separation. To accomplish this, the chamber is filled about 1 inch from the bottom with the appropriate solvent (depends on what you are separating). A large sheet of chromatography paper (Whatman 3MM Chr) is added to the tank. This should be cut to fit in the tank by curving around the sides of the tank (See Picture). Vacuum grease is used to seal the chamber (See Picture). This vacuum chamber should be set up the night before use.
2.) Standard Whatman Thin Layer Plates are used (LK6D Silica Gel 60A) for this procedure. They must be washed with acetone in a regular solvent tank for 1 hour and then dried in a fume hood before use (See Picture).
3.) Individual plates are marked with a pencil 20mm from the top of the loading area. This marks where the lipid will be applied (See Picture). When the elution reaches 10mm from the top of the plate, the plate is removed from the chamber and another mark is made at this point.
4.) Lipid is added to the individual lanes of the TLC plates using capillary pipettes (Fisherbrand; 20microliters). 20-50 microliters of lipid is added (See Picture). After blotting, the plate is placed in the solvent chamber with appropriate solvent (See Picture). The elution usually reaches the upper marking of the plate in about 1 hour and then the plate is removed.
5.) After the plate is allowed to dry (1-2 hours), the TLC plate is analzed. This may involve sprays, stains,14C, 3H, fluorescence, etc. In this case 14C and 3H were used. The lipids on the left side of this plate had been tagged with 3H. Therefore, these lanes are sprayed with an autoradiographic enhancer specific for 3H (NEN). In a fume hood, the TLC plate is sprayed (See Picture), allowed to dry for 10 minutes, rotated 90o and then sprayed again.
6.) Radioactive dye is added to the 4 corners of the TLC plate (See Picture) so that it can be lined up with the film after developing (see below).
7.) The TLC plate is wrapped in plastic wrap, taped and placed in an autoradiography cassette (Biotech) (See Picture). In a dark room, Kodak Biomax MR film is added to the cassette. The cassette is wrapped in aluminum foil and placed in a -80o freezer for 72 hours (See Picture).
*NOTE: An alternative to Step #7 may be the use of a phosphoimager especially for 32P fluorescence. This allows for quantification by densitometry without cutting the bands out.
8.) After 72 hours, the cassette is removed from the -80o freezer and allowed to warm to room temperature (approximately 2 hours).
9.) After developing, (See Picture) the film can then be analyzed qualitatively for specific bands. To quantify the spcific bands seen, the TLC plate is matched up with the film (corner markers) to mark the bands on the TLC plate. A large bore needle is used to mark the bands See Picture below.
10.) For this step gloves and a mask must be worn. After the bands are marked, the TLC plate is misted with water to keep the dust to a mininum. The individual bands are then removed with a razor blade (See Picture) and placed in appropriately labeled scintillation vials (See Picture). These vials are agitated using a Vortex-Genie and allowed to stand overnight. They are then placed in a scintillation counter (See Picture). The counts per minute obtained allow for quantitative comparison between the different bands.
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References:
Dr. Bill Stillway (stillway@musc.edu), Ben Pettus (pettusbj@musc.edu).
Jenkins GM and Hannun YA. Sphingolipids as messengers of cell death. In Studzinski GP (Ed.), Apoptosis: A Practical Approach. New York: Oxford University Press, 1999, p. 105-123.
Kates, Morris. Techniques of Lipidology: Isolation, Analysis, and Identification of Lipids. In Burden RH, van Kippenburg PH (Eds.), Laboratory Techniques in Biochemistry and Molecular Biology. New York: Elesvier, p. 232-254.
