Kansas Lipidomics Research Center

Lipid Extraction Method for Arabidopsis (and other) Leaves (Method 1)

This procedure is generalized. Use of smaller samples is possible. This procedure can be adapted for other plant tissues, including roots, stems, flowers, and siliques. Please contact Mary Roth at mrroth@ksu.edu (please include a subject line) to inquire.

A .pdf of this method, including shipping directions, can be found here.

  1. Take 1 to 8 leaves (or up to 3 whole little plants- see NOTE #2); quickly immerse in 3 ml 75ºC (preheated) isopropanol with 0.01% BHT (butylated hydroxytoluene, e.g., Sigma B1378) and continue to heat for 15 min. Use a 50 ml (25 x 150 mm) glass tube with a Teflon-lined screw cap. NOTE #1: It is extremely important that the plants be extracted immediately after sampling and that the isopropanol be preheated. Plants have very active phospholipase D, which is activated upon wounding; failure to place the sampled tissue quickly into hot isopropanol will result in generation of phosphatidic acid. NOTE #2: To do the analysis, only a fraction of an Arabidopsis leaf is needed. Use of more tissue reduces variability among samples; use of smaller amounts of tissue can provide information about individual plants and plant tissues. Dry weights of 5 to 30 mg, measured as in step 6, are recommended.

  2. Add 1.5 ml chloroform and 0.6 ml water, vortex; then agitate (shaking incubator) at room temperature for 1 hour. Transfer (long, glass Pasteur pipettes) lipid extracts to glass tubes with Teflon-lined screw-caps.

  3. Add 4 ml chloroform/methanol (2:1) with 0.01% BHT; shake 30 min. Repeat this extraction procedure on all samples until the leaves of every sample become white, but be sure to extract each sample the same number of times. (Use one pipette in each sample for all extractions, leaving them in the removed extract while extracting the remaining materials.) It's OK to leave the tubes shaking for somewhat longer than 30 min on later extractions, and leaving one of the extractions shaking overnight is a good idea for difficult-to-extract tissues. Usually you will need about 5 extractions, including the one with the isopropanol.

  4. Optional back-washes: Add 1 ml 1 M KCl to the combined extract, vortex or shake, centrifuge, discard upper phase. Add 2 ml water, vortex or shake, centrifuge, discard upper phase. These backwashes will yield a cleaner lipid sample, but small amounts of the more polar lipids, such as lysolipids, will be lost.

  5. Fill tubes containing lipid extract with nitrogen, store at -20°C (freezer). When you are ready to prepare for shipping, evaporate solvent completely under nitrogen gas or in a speedvac, and redissolve in about 1.0 ml chloroform. Transfer to 2.0 ml clear glass vial with Teflon-lined screw cap (for example: clear glass; 2 mL; solid PTFE-lined cap; 1/2 dr., Fisher Scientific catalog #03-391-7A, Thermo Scientific No.: B7800-1). Evaporate solvent from the samples in the 2-ml vials before shipping. ALWAYS let us know before you send your samples: mrroth@ksu.edu

  6. Dry extracted leaves at 105°C oven overnight; weigh for "dry" weights preferably using a balance that weighs (in grams) to 6 decimal places (i.e., micrograms).

Video demonstration of Method 1:

Click here to watch at YouTube (full screen available)

Lipid Extraction Method for Leaves (Method 2, tested on Arabidopsis and sorghum)

A more rapid method of lipid extraction was recently described by Shiva et al. (2018): https://plantmethods.biomedcentral.com/articles/10.1186/s13007-018-0282-y

This approach gets a good extraction from leaves in a single step. If following Method 2, after separating the plant materials from the lipid extract, follow steps #5 and #6 in the above protocol.

Video demonstration of Method 2:

Click here to watch at YouTube (full screen available)

Lipid Extraction Method for Leaves (Method 3 - if you need GIPC analysis, in addition to that of other lipids)

If you want to include glycosylinositolphosphoceramide (GIPC) in your extract, you need to do additional extractions of your leaves with "Solvent H".

Solvent H from Markham et al. (2006, http://www.jbc.org/content/281/32/22684.full.html):
Isopropanol/hexane/water 55:20:25, upper phase discarded
440 ml 2-propanol w/0.01% BHT (0.04g)
160 ml hexane (HPLC grade)
200 ml water

Mix these together (stirring vigorously), and remove the upper phase for discard. Lower phase should be clear and is Solvent H.

  1. Complete either steps 1-3 in Method 1 or the Method 2 one-step method.

  2. To the remaining leaf tissue, add 4 ml “Solvent H”. Put tube on heating block at 60°C for 15 min. Remove solvent and combine with the chloroform/methanol lipids extracted previously.

  3. Repeat three more times for a total of 4 “Solvent H” extractions. The total volume of all extracted lipids plus solvents is about 37 ml (if you used Method 1).

  4. Fill tubes with nitrogen, store at -20°C (freezer). When you are ready to prepare for shipping, evaporate completely under nitrogen gas or a speedvac, at less than 40ºC, and redissolve in about 1.0 ml chloroform. Transfer to 2.0 ml clear glass vial with Teflon-lined screw cap (for example: clear glass; 2 ml; solid PTFE-lined cap; 1/2 dr., Fisher Scientific catalog #03-391-7A, Thermo Scientific No.: B7800-1). Evaporate solvent from the samples in the 2-ml vials before shipping. ALWAYS let us know before you send your samples: mrroth@ksu.edu

  5. Leaves extracted with Solvent H will stick to the glass when dry, unless a bit of chloroform is added and the leaves are pushed into a ball. A spatula works well for forming the ball. Dry extracted leaves at 105°C in an oven overnight, and weigh to 6 decimal places (0.000000g) for dry extracted tissue weight. (Just dump the dried tissue into a weigh boat.)