Thin layer chromatography (Thiele Lab)

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Thin layer chromatography (Thiele Lab)

This procedure uses two sequential chromatographic steps on one TLC plate to give excellent resolution of both polar and neutral lipids.

Step 1:
Take a 20x20 cm silica gel TLC plate (Silica gel 60 Merck 1.05721.001, without UV indicator) With a soft pencil, draw a line at a distance of 1.5 cm of one of the edges. On this line, mark at equal distances the application zones for your samples and a set of standard lipids for identification (e.g. SM, PC, PE, PI, cholesterol, cholesterol ester, triglyceride).

Step 2:
Apply your samples, dissolved in chloroform/methanol 2/1 to the plate using a glass capillary. Take care to apply the sample as a narrow line, not as a large round spot. If you have 30 µl of sample, you will have to load it in 3-5 rounds on top of each other. Let it dry for 1-2 min between each round of loading.
Capacity: On one lane, load the extracted lipids equivalent to 50-100% of a 10 cm dish of cell culture. For very lipid-rich cell lines (e.g. differentiated 3T3-L1) take less.

Step 3:
Dry the plate under a gentle stream of air at room temperature, until it does not show wet spots any more if you inspect it in trans-illumination. This is crucial, since water on the plate will result in heavily distorted bands.

Step 4:
Equilibrate two TLC chambers with solvent mix I and II. Use filter paper that you place to the front and back sides of the chamber to support complete equilibration.

solvent mix I: CHCl3/triethylamine/ethanol/H20  35/35/40/9

solvent mix II: Isohexane/ethyl acetate 5/1

Step 5:
Develop the plate in solvent I
Let the solvent run until the front is about 12-14 cm above the start line (about 45 min). Take the plate out, mark the front at the edges with a soft pencil and dry it with a hair dryer (warm, not hot).

Step 6:
Develop the plate in solvent II, until the front is 1 cm below the upper edge of the plate (about 30 min).

Some general remarks to the choice of the solvent systems: (i) On a silica plate, non-polar substances run faster than polar substances. (ii) The more polar the solvent mix, the faster is the running behaviour of the substances. (iii) Each solvent mix will in general only resolve a certain part of a complex mix of substances. Some hydrophobic substances will run with the solvent front while some hydrophilic substances will stay on the start line.
In our case, resolution with mix I includes lipids running from lyso-PC, SM, all phospholipids, free fatty acids, monoglycerides until cholesterol. Complex glycolipids will stay at the start line, cholesterol ester, triglycerides and other very hydrophobic lipids will run unresolved with the solvent front. To resolve the complex glycolipids, more hydrophilic solvent mixtures (e.g. CHCl3/MeOH/0.25% KCl in H20 65/35/8) would be used. The second run with mix II will resolve hydrophobic neutral lipids whereas the polar lipids do not move in this mix and stay at their initial positions. If you only want to resolve neutral lipids, and do not care about the polar lipids, you may use a single mix such as Hexane/Ether/Acetic acid 90/10/1.


Detection
To visualize the lipids by sulphuric acid staining:

Spray the plate with 20% sulfuric acid (lab coat, gloves, protective glasses!!!) and heat it to 150-200 degree C.

Watch the staining process from time to time. First, two wine-red bands in the upper third will appear. This is cholesterol and cholesterol ester; this staining is very sensitive and diagnostic. Upon prolonged heating, most other lipids will give yellow-green-brown bands. Take the plates out at this moment. All bands would turn black upon long-term heating. but the colors contain some useful information. Most of this staining depends on the presence of double bonds or sugar moieties in the lipids; therefore, strongly unsaturated lipids will be over-represented while (the very rare) disaturated phospholipids stain poorly (for example, di-palmitoyl PC fails to stain at all).

Identify the lipid bands by comparison with the standards.


Important comment on chloroform: Chloroform can chemically decompose, driven by light and oxygen, into highly reactive compounds. In particular, HCl, phosgen and chlorine will be formed. In order to avoid this, chloroform is usually doted with a stabilizer such as ethanol or amylene. Particularly, amylene-stabilzed chloroform tends to cause problems. It is highly recommended to use ethanol-stabilized chloroform with a certificate for absence of phosgene and HCl, such as the one offered by Fluka (No. 25690)



Literature:

For TLCs run with this system, see 1,2

1.    Kuerschner, L., Ejsing, C.S., Ekroos, K., Shevchenko, A., Anderson, K.I. & Thiele, C. Polyene-lipids: A new tool to image lipids. Nat Methods 2, 39-45 (2005)
2.    Kuerschner, L., Moessinger, C. & Thiele, C. Imaging of Lipid Biosynthesis: How a Neutral Lipid Enters Lipid Droplets. Traffic 9, 338-352 (2008)


More information on TLC, particularly on special TLC systems for certain lipid subfractions, and specific stains for lipid classes, you will find in the old but useful small red book:

Morris Kates, Techniques of Lipidology, New York: Elsevier 1986



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