Lipid Isolation from Plants

Abstract

Analysis of plant lipids provides insights into a range of biological processes, from photosynthetic membrane function to oil seed engineering. Many lipid extraction protocols are tailored to fit a specific lipid class. Here we describe a procedure for extraction of glycerolipids from vegetative tissue. This procedure is designed for 1 gram of tissue per sample but maybe scaled for larger samples.

1. Introduction

Plant lipids are essential components of every tissue type and as such, accurate measurements of their composition are important to a thorough understanding of plant biology. Lipid analysis can be a routine procedure which greatly amplifies a plant scientist’s toolbox. Critical to this analysis is understanding that plant tissues, particularly leaves, are replete with a range of lipases that complicate lipid extraction. Additionally, the fatty acid substituents of lipids can oxidize unless proper care is taken to protect polyunsaturated fatty acids from damage [1].

Phospholipases present in vegetative tissues can rapidly alter the lipid composition, generating artifacts which can be difficult to identify. Products of lipase activity may be misidentified as other lipids of interest, leading to embarrassing errors [2]. The problem begins with tissue disruption, releasing phospholipases which frequently remain active in the organic solvents used for lipid extractions, even retaining activity at low temperatures ( 16 °C) in some cases [3, 4]. Detection of intermediates in lipid synthesis such as lysophospholipids, free fatty acids, and phosphatidic acid as large fractions of the lipid sample may indicate phospholipase activity [1]. Inactivation of this potential lipase activity is essential for successful plant lipid extraction. A straightforward way to achieve this is to use a solvent system containing formic or acetic acid at cold temperature. Boiling the sample in isopropanol with 0.01% butylated hydroxytoluene (BHT) as antioxidant before proceeding with the lipid extraction also reduces artifact production [5].

Lipid extractions typically use chloroform, methanol, and water, in proportions that produce a monophasic mix. A classic example of this is the Bligh and Dyer method designed for the high water content of fish tissue [6]. It is important to note that a protocol designed for plant vegetative tissue may not need the addition of water, due to the high-water content of leaves, while lipid extraction from dry seeds will require added water. Generating a monophasic mix permits efficient lipid extraction from the tissue. This is followed by addition of salts to obtain a phase separation leading to partition of the lipids. Nonlipid components segregate into the water-rich layer while most lipids partition to the chloroform layer. However, highly polar lipids may partition partially into the aqueous phase, in an equilibrium responsive to negative charges on the lipid. The addition of acids such as phosphoric acid produces mildly acidic conditions that protonate charged lipids and neutralize their charge, so that they partition more efficiently into the chloroform layer (Fig. 1a). Chloroform–methanol–water procedures readily extract glycerolipids, but some sphingolipids are not readily extracted, and a more involved extraction than discussed here is required [7].

Fig. 1.

Lipid isolation and quantification by GC separation of FAMEs. (a) Phase separation of an extract from an Arabidopsis thaliana leaf sample provides for partitioning of the lipids into the lower chloroform layer which is green due to the presence of chlorophyll. (b) Separation of FAMEs derived from Arabidopsis leaf lipids by GC with 17:0 as internal standard allows quantification of the total glycerolipids in the sample

Here we describe a simple method for extracting glycerolipids from vegetative plant tissue. This procedure limits oxidation of fatty acids by keeping temperatures low, and inhibits lipase activity with acetic acid. Like both the Folch [8], and Bligh and Dyer [6] methods, we use chloroform, methanol, and water (present in the vegetative tissue) along with acetic acid to generate a monophasic solution. We induce phase a separation by adding aqueous KCl and phosphoric acid. Lipids partition to the lower chloroform layer. Neutral and most polar lipids readily partition into the chloroform layer, but highly polar lipids such has lysophosphatidic acid maybe partially soluble in the aqueous methanol and water layer. Quantitative extraction of these lipids is achieved by back extraction with chloroform. Other lipids, particularly some intermediates such as acyl-ACPs and acyl-CoAs, are soluble in the water–methanol layer and may bind to protein at the water–chloroform interface [9, 10].

Estimating the quantity of glycerolipids from a sample is accomplished by derivatizing the substituent fatty acids to methyl esters. The fatty acid methyl esters (FAME) can be separated and analyzed by gas chromatography with a flame ionization detector (GC-FID). Use of an internal standard when derivatizing fatty acids esterified to glycerolipids allows for easy quantification of FAMEs. An internal triacylglycerol standard such as triheptadecanoylglycerol (17:0-TAG) normalizes for the efficiency of fatty acid derivatization. Detection of FAMEs by FID provides a nearly linear response to the carbons in the acyl chains, allowing for reliable quantification of the sample (Fig. 1b).

2. Materials

2.1. Preparing Solvents

1. Fume hood.

2. Glass graduated cylinder (1 L).

3. Storage bottles with solvent-resistant polypropylene caps.

4. Chloroform (HPLC grade).

5. Methanol (HPLC grade).

6. Glacial Acetic acid.

7. Phosphoric acid.

8. Potassium chloride.

9. Toluene (HPLC grade).

10. Butylated hydroxy toluene (BHT).

11. Sulfuric acid.

12. 2 L Erlenmeyer flask.

2.2. Harvesting Tissue and Storage

1. 50 mL Falcon tubes (e.g., Fisher Scientific, Cat # 14–432-22).

2. Two tube racks accommodating 50 mL Falcon tubes.

3. Liquid nitrogen.

4. Vacuum insulated Dewar flask for liquid nitrogen.

5. Two insulating foam boxes.

6. Sharp scissors.

7. Balance with sufficient precision for your samples.

8. −80 °C freezer.

2.3. Tissue Homogenization

1. Mortar and pestle.

2. 30 mL solvent-resistant fluorinated ethylene/propylene polymer Nalgene tubes.

3. 10 mL glass pipettes and safety bulb or pipette plunger.

4. Liquid nitrogen.

5. Solvent mixture chloroform–methanol–glacial acetic acid (10:10:1, v/v/v) (12 mL per sample).

2.4. Lipid Extraction

1. Refrigerated centrifuge that can accommodate solvent-resistant tubes.

2. 25 mL borosilicate tubes.

3. Teflon lined caps for 25 mL glass tubes.

4. Glass Pasteur pipettes.

5. Pipette bulb for glass Pasteur pipettes.

6. Vortex mixer.

7. Clinical centrifuge suitable for 25 mL glass tubes.

8. Organic extraction solution chloroform–methanol–acetic acid (5:5:1, v/v/v) (5 mL per 1 g tissue sample).

9. Extraction buffer 1 M KCl, 0.2 M H₃PO₄ (6 mL per 1 g tissue sample).

10. Gas evaporation manifold equipped with a heating block able to heat samples at 40 °C.

11. Nitrogen gas.

2.5. Quantification of Total Lipids

1. Gas chromatograph with spilt/splitless injector and flame ionization detection (see Note 1).

2. Carbowax capillary GC column (e.g., SUPELCOWAX ® 10, 0.53 mm, 15 m, Sigma-Aldrich).

3. Triheptadecanoylglycerol (17:0-TAG) standard.

4. Toluene (HPLC grade).

5. Butylated hydroxy toluene (BHT).

6. Pipettes (P200) or Drummond Digital Microdispenser (see Note 2).

7. 8 mL glass tubes.

8. Teflon-lined caps for glass tubes.

9. Hot water bath set to 85 °C.

10. Hexanes (mixture of hexane isomers, e.g., Sigma-Aldrich HX0296) or n-hexane.

11. GC vials (1 mL) with caps.

12. GC vial inserts (300 μL).

3. Methods

3.1. Preparation of Solvents (Protocol Designed for Samples of 1 g or Less)

1. The solvent mixtures should be prepared 1 day before extracting lipids. Because the mixtures contain chloroform, a known carcinogen, use a fume hood and wear gloves, goggles and a lab coat when working with chloroform.

2. Prepare chloroform–methanol–glacial acetic acid (10:10:1, v/v/v) (see Note 3).

   Prepare sufficient volume for 12 mL per sample with some to spare.

   Using a glass graduated cylinder and working in a fume hood, mix the appropriate volumes of chloroform, methanol and acetic acid together into a borosilicate bottle, then cap the bottle and store at −20 °C.

3. Using a glass graduated cylinder and working in a fume hood, prepare chloroform–methanol–acetic acid (5:5:1, v/v/v).

   Prepare sufficient volume to use 5 mL per sample, with some to spare.

4. Prepare 1 M KCl, 0.2 M H₃PO₄ (6 mL per g of sample). For 1 L: Add to 74.6 g of KCl to 500 mL of deionized water in a 2 L Erlenmeyer flask.

   Add a magnetic stir bar and place on a stir plate, gently stirring the sample.

   Using a graduated cylinder measure 35.3 mL of 85% (w/v) phosphoric acid.

   Add the phosphoric acid to the Erlenmeyer flask containing KCl in 500 mL of deionized water.

   When the KCl has completely dissolved adjust volume to 1 L in a glass graduated cylinder and store in a tightly capped glass bottle.

5. Prepare 0.005% BHT in toluene (see Note 4).

   Measure 500 mL of toluene with a glass graduated cylinder.

   Transfer to a glass bottle with solvent-resistant cap and add 25 mg of BHT.

   Allow BHT to dissolve in toluene.

6. Prepare 2.5% (v/v) sulfuric acid in methanol.

   Appropriate attire for this exothermic reaction includes lab coat, googles and gloves.

   In a fume hood very slowly add 25 mL sulfuric acid to 975 mL of methanol.

   Danger: Failure to add the sulfuric acid slowly can result in rapid boiling and splashing of sulfuric acid, potentially burning exposed tissue (see Note 5).

   Store the reagent in a capped glass bottle at room temperature.

7. Prepare the internal standard.

   Dissolve 100 mg of 17:0-TAG in 200 mL of toluene with 0.005% (w/v) BHT.

   This provides a final concentration of 0.5 μg standard per μL.

   Store in Teflon-capped glass bottle at 20 °C until use. Warm to room temperature prior to use.

3.2. Harvesting Tissue for Storage

1. Appropriate attire includes lab coat, googles and gloves. Liquid nitrogen used in this process is dangerous and will burn skin on contact.

2. Label Falcon tubes.

3. Put a tube rack in an insulating foam box and place labeled 50 mL Falcon tubes into the tube rack.

4. Pour approximately 4 cm of liquid nitrogen into the box.

   Liquid nitrogen can burn skin. Carefully pour the liquid nitrogen, avoiding skin contact.

5. Fill the Falcon tubes with 2 to 3 cm of liquid nitrogen; do not cap (see Note 6).

6. Place a second tube rack in a second foam box without liquid nitrogen.

7. Cut approximately 1 g of tissue per sample and quickly weigh it using the balance.

8. Transfer the tissue from the scale to a Falcon tube containing liquid nitrogen.

9. Place the tube containing the weighed sample into the second box, not immersed in liquid nitrogen.

10. Allow the liquid nitrogen in the tube to completely evaporate, then immediately cap each tube while the sample is still frozen and place it in the tube rack immersed in liquid nitrogen. Proceed to the next sample.

11. Store samples at −80 °C until extraction.

3.3. Tissue Homogenization

1. Label 30 mL solvent-resistant Nalgene tubes and place on ice.

2. Place the bottle of chloroform–methanol–glacial acetic acid (10:10:1, v/v/v) on ice in the fume hood.

3. Chill the mortar and pestle by filling the mortar with liquid nitrogen, placing the pestle in the mortar (see Note 7).

4. When the liquid nitrogen stops boiling the mortar and pestle are ready to use.

5. Transfer the first sample from the Falcon tube to the liquid nitrogen in the mortar.

6. Homogenize the frozen sample with the pestle, initially crushing the sample gently into smaller pieces, then grinding with the pestle until the sample is powdered (see Note 8).

   It is best to rapidly grind the tissue when much of the liquid nitrogen has evaporated, leaving tissue with the appearance of snow.

7. Transfer 5 mL of chloroform–methanol–acetic acid (10:10:1, v/v/v) to the mortar with a 10 mL glass pipette and manual pipette pump (see Note 9). Do not mouth pipette.

8. Resuspend the tissue in the solvent mixture and pour into a labeled 30 mL solvent-resistant fluorinated ethylene propylene tube.

9. Rinse the mortar and pestle with an additional 5 mL of chloroform–methanol–acetic acid (10:10:1, v/v/v) and combine the wash with the sample in the solvent-resistant tube.

10. Rinse the mortar and pestle again with 2 mL of chloroform– methanol–acetic acid (10:10:1, v/v/v) and combine the material into the solvent-resistant tube.

11. Wipe the mortar and pestle with a paper towel and cool again with liquid nitrogen before proceeding to the next sample.

12. Store the samples at 20 °C at least 4 h (or overnight) to allow for a complete extraction of lipids.

3.4. Lipid Extraction

1. Cool the centrifuge and rotor to 4 °C.

2. Place the bottle of chloroform–methanol–acetic acid (5:5:1, v/v/v) on ice in the fume hood.

3. Label a second set of 30 mL solvent-resistant tubes and place on ice.

4. Label a set of 25 mL glass tubes.

5. Remove the samples from the −20 °C freezer and centrifuge at 4000 × g at 4 °C for 5 min to pellet the debris.

6. Taking care not to disturb the pelleted debris, transfer the supernatant to a new labeled tube using a glass Pasteur pipette (see Note 10).

7. Wash the pellet by adding 5 mL of cold chloroform–methanol– acetic (5:5:1, v/v/v) acid to the 30 mL tube containing the pellet, cap and mix thoroughly with a vortex mixer.

8. Centrifuge at 4000 × g and 4 °C for 5 min.

9. Combine the wash supernatant with the supernatant from the sample, taking care to only transfer the supernatant to the new tube. Discard the pellet.

10. Create a phase separation by the addition of 6 mL 1 M KCl in 0.2 M H₃PO₄ to the new tube (see Fig. 1a). Cap and vortex the tubes.

    Centrifuge at 4000 g and 4 °C for 5 min to aid the phase separation.

    Transfer the bottom chloroform layer to a clean labeled 25 mL glass tube, being careful to take only the chloroform layer.

    Back-extract the aqueous layer with an additional 5 mL of chloroform.

    Cap and vortex the tube.

    Centrifuge and combine the lower chloroform layer with the first extraction, being careful to take only the chloroform layer.

11. Evaporate the chloroform under nitrogen in the gas evaporation manifold, with the heating block set to 40 °C.

12. Resuspend in 500 μL toluene, 0.005% BHT (see Note 11). Close with a Teflon-lined cap.

    Gently vortex the sample to dissolve any lipids on the tube wall.

13. For storage keep the sample at 20 °C where it will be stable for at least 6 months.

3.5. Quantification of Total Lipids by GC Analysis of Fatty Acid Methyl Esters

1. Warm the 17:0-TAG standard in toluene to room temperature (see Note 12).

2. Using either a P200 or 100 μL Drummond microdispenser place 100 μL of internal standard into an 8 mL glass tube as a standard-only control.

3. Using either a P200 or 100 μL Drummond microdispenser take a 50 μL aliquot of lipid extract and place in a labeled 8 mL glass tube, add 100 μL (50 μg) of 17:0-TAG internal standard.

4. Using 10 mL glass pipettes, add 1.5 mL of 2.5% sulfuric acid in methanol to each of the 8 mL glass tubes (see Note 13).

5. Close tightly with a Teflon-lined cap.

6. Check that the water bath is at 85 °C and place the rack of samples in it. After 5 min check tubes to make sure the caps are tight, and no solvent is evaporating. If solvent appears to be evaporating replace evaporated solvent and recap the tube before returning it to the bath.

7. Continue heating at 85 °C for 1 h in the water bath.

8. Cool to room temperature.

9. Add 200 μL of hexanes and 1 mL of water and recap the tubes.

10. Shake or vortex the tubes for 30 s, then centrifuge at less than 1000 × g for 2 min (see Note 14).

11. Label 1 mL GC vials.

12. Using either a P200 or 100 μL Drummond microdispenser remove 70–100 μL of the hexane (top) layer to a glass insert, place in the labeled GC vial, and cap.

13. Prepare the GC for analysis.

14. FAMEs can be separated by using a carbowax capillary column (e.g., SUPELCOWAX® 10, 0.53 mm, 15 m) (Fig. 1b) (see Note 15).

15. An example of GC conditions is as follows: 1 μL sample injected.

    Start temperature 190 °C for 1 min.

    Temperature ramp increases by 7 °C per min to 250 °C.

16. Use software of choice to output peak areas.

17. Quantify total lipid based on the area of the 17:0 standard peak (see Note 16).

    μg/area = 50 μg/area of 17:0.

    Total μg in sample = (sum of all peak areas except 17:0 peak area) × μg/area.

    μg/μL fatty acid in lipid extraction(total μg in sample/ 50 μL) (see Note 17).