Determination of Total Lipid and Lipid Classes in Marine Samples

Summary

This protocol is for the determination of lipids in seawater and biological specimens. Lipids in filtrates are extracted with chloroform or mixtures of chloroform and methanol in the case of solids. Lipid classes are measured by rod thin-layer chromatography with flame ionization detection and their sum gives the total lipid content.

Abstract

Lipids are largely composed of carbon and hydrogen and, therefore, provide a greater specific energy than other organic macromolecules in the sea. Being carbon- and hydrogen-rich they are also hydrophobic and can act as a solvent and absorption carrier for organic contaminants and thus can be drivers of pollutant bioaccumulation in marine ecosystems. Their hydrophobic nature facilitates their isolation from seawater or biological specimens: marine lipid analysis begins with sampling and then extraction in non-polar organic solvents, providing a convenient method for their separation from other substances in an aquatic matrix.

If seawater has been sampled, the first step usually involves separation into operationally defined 'dissolved' and 'particulate' factions by filtration. Samples are collected and lipids isolated from the sample matrix typically with chloroform for truly dissolved matter and colloids, and with mixtures of chloroform and methanol for solids and biological specimens. Such extracts may contain several classes from biogenic and anthropogenic sources. At this time, total lipids and lipid classes may be determined. Total lipid can be measured by summing individually determined lipid classes which customarily have been chromatographically separated. Thin-layer chromatography (TLC) with flame ionization detection (FID) is regularly used for the quantitative analysis of lipids from marine samples. TLC-FID furnishes synoptic lipid class information and, by summing classes, a total lipid measurement.

Lipid class information is especially useful when combined with measurements of individual components e.g., fatty acids and/or sterols, after their release from lipid extracts. The wide variety of lipid structures and functions means they are used broadly in ecological and biogeochemical research assessing ecosystem health and the degree of influence by anthropogenic impacts. They have been employed to measure substances of dietary value to marine fauna (e.g., aquafeeds and/or prey), and as an indicator of water quality (e.g., hydrocarbons).

Introduction

The methods described here concern substances that are defined operationally as marine lipids. This definition is based on their amenability to liquid-liquid extraction in non-polar organic solvents, and it provides a convenient method for their separation from other substances in an aquatic matrix. Their hydrophobic nature facilitates their isolation from seawater or biological specimens, as well as their enrichment, and the removal of salts and proteins.

The measurement of lipid content and its composition in marine organisms has been of great interest in food web ecology, aquaculture nutrition, and food science for decades. Lipids are universal components in living organisms, acting as essential molecules in cell membranes, as major sources of bioavailable energy, providing thermal insulation and buoyancy, and serving as signaling molecules. Although procedures for lipid determination in other fields have been described well, their use with marine samples commonly necessitates modification to adapt to field conditions as well as to sample type¹.

For seawater samples, the first step usually requires separation into the operationally defined 'dissolved' and 'particulate' fractions, normally by filtration (Protocol step 1). The particulate fraction is what is retained by the filter, and size of the pores is important in defining the cut-off². Often when we are sampling particulate matter, we would like to relate lipid concentrations to total mass concentrations, in which case a separate, smaller, sample (e.g., 10 mL) has to be taken for this purpose (Protocol step 1, note). To get an accurate mass determination it is important to add ammonium formate (35 g/L) at the end of the filtration.

The seawater filtrate from the larger sample should amount to between 250 mL and 1 L depending on sample type and is subjected to liquid-liquid extraction in a separatory funnel (Protocol step 2). The hydrophobic nature of lipids means they can be separated from other compounds by extraction in a nonpolar solvent such as chloroform. A two-layer system is created where lipids partition into the organic layer while water soluble components stay in the aqueous layer.

Particulate samples on a filter, or biological specimens are extracted with a modified Folch et al. extraction³, also involving chloroform (Protocol step 3). Again, an organic/aqueous system is created in which lipids partition into the organic phase, while water soluble molecules remain in the aqueous phase, and proteins are precipitated. In fact, for solids, most laboratories use some variation of the Folch et al. extraction³ procedure involving chloroform and methanol. For filters, the first step is to homogenize in 2 mL of chloroform and 1 mL of methanol.

During extraction, care should be taken to protect lipids from chemical or enzymatic modification, by keeping samples and solvents on ice to reduce ester bond hydrolysis or carbon-carbon double bond oxidation. Tissues and cell lipids are quite well protected by natural antioxidants and by compartmentalization⁴; however, following the homogenization of samples, cell contents are combined rendering lipids more disposed to alteration, chemically or enzymatically. Some lipids, such as most sterols, are very stable, while others, such as those containing polyunsaturated fatty acids, are more susceptible to chemical oxidation. Others, such as sterols with conjugated double bonds, are prone to oxidation catalyzed by light⁵. Following extractions, lipids are much more susceptible to chemical oxidation, and samples should be stored under an inert gas such as nitrogen. A gentle stream of nitrogen would also be used to concentrate extracts.

After concentration, lipids would then normally be quantified in bulk as they are an important component of marine ecosystems providing a high concentration of energy, more than twice the kJ/g of carbohydrates and proteins. Invariably they would next be quantified as individual components: the comprehensive analysis of lipids generally involves separation into simpler categories, according to their chemical nature. Thus, a full analysis involves measuring total lipids, lipid classes and individual compounds.

Total lipid can be determined by taking the sum of individually measured lipid classes separated by chromatography⁶. A marine lipid extract may contain more than a dozen classes from biogenic and anthropogenic sources. The wide variety of lipid structures means much information can be gained by determining individual groupings of structures. Lipid classes individually, or in certain groups, have been used to signal presence of certain types of organisms, as well as their physiological status and activity². They have also been used as an indicator of the origins of organic material, including dissolved organic matter (DOM) as well as hydrophobic contaminants.

Triacylglycerols, phospholipids and sterols are among the more important biogenic lipid classes. The first two are biochemically related as they possess a glycerol backbone to which two or three fatty acids are esterified (Figure 1). Triacylglycerols, together with wax esters are very important storage substances, while other fatty acid-containing lipid classes such as diacylglycerols, free fatty acids, and monoacylglycerols are generally minor constituents. Free fatty acids are present at lower concentrations in living organisms, as the unsaturated ones can be toxic⁷. Sterols (both in their free and esterified forms) and fatty alcohols are also included among the less polar lipids, while glycolipids and phospholipids are polar lipids. Polar lipids have a hydrophilic group, which allows for the formation of lipid bilayers found in cell membranes. Free sterols are also membrane structural components, and when taken in ratio to triacylglycerols they provide a condition or nutritional index (TAG : ST) which has been widely used⁸. When taken in ratio to phospholipids (ST : PL) they can be used to indicate plant sensitivity to salt: higher values maintain structural integrity and decrease membrane permeability⁹. The inverse of this ratio (PL : ST) has been studied in bivalve tissues during temperature adaptation¹⁰.

Marine lipid classes can be separated by thin-layer chromatography (TLC) on silica gel coated rods (Protocol step 4) and then detected and quantified by flame ionization detection (FID) in an automatic FID scanner. TLC/FID has become routinely used for marine samples as it rapidly furnishes synoptic lipid class data from small samples, and by taking the sum of all classes, a value for total lipids. TLC/FID has been subjected to a quality-assurance (QA) assessment and was found to meet standards required for consistent external calibration, low blanks, and precise replicate analysis¹¹. Coefficients of variation (CV) or relative standard deviations are around 10%, and FID scanner total lipid data are normally around 90% of those obtained by gravimetric and other methods². Gravimetry gives higher total lipids likely because the FID scanner measures only non-volatile compounds, and also as a result of possible inclusion of non-lipid material in gravimetric measurements.

The information provided by lipid class analysis is especially useful when combined with determinations of fatty acids as individuals, or sterols, or the two in combination. The first step towards these analyses involves the release of all component fatty acids together with sterols in the lipid extracts (Protocol step 5). The wide variety of lipid structures and functions means they have seen broad use in ecological and biogeochemical studies assessing ecosystem health and the extent to which they have been influenced by anthropogenic and terrestrial inputs. They have been used to measure biosynthesis of substances of dietary value to marine fauna as well as to indicate the quality of water samples. Measuring lipids in sediment core samples helps show the sensitivity of sediments to changes in human land use near the land-sea margin.

The primary tool for identifying and quantifying individual lipid compounds has traditionally been gas chromatography (GC) with FID. Before analysis however, these compounds are made more volatile by derivatization. Fatty acids are released in the presence of an acidic catalyst (H₂SO₄) from acyl lipid classes (Figure 1). In organic chemistry, the acyl group (R-C=O) is usually derived from a carboxylic acid (R-COOH). They are then re-esterified to fatty acid methyl esters (FAME) which gives better separations on GC columns (Protocol step 5).

Protocol

NOTE: To clean glassware, instruments and filters for lipid analyses, wash them 3 times with methanol followed by 3 washes with chloroform, or heat them to 450°C for at least 8 hours.

1. Filtration procedure for seawater dissolved and particulate lipids

NOTE: The particular fraction of interest is operationally defined by the filtration procedure. In this case the pore size is 1.2 µm.

1. Set up the filtration manifold without a filter and rinse the setup with filtered seawater.
2. Using clean forceps place a 47 mm glass fiber (GF/C) filter, that has been washed, into the clean filtration system.
3. Take the sample and gently swirl to resuspend any material that may have settled on the bottom of the collection container. Accurately measure out a known volume, in this case 1 L, and filter this through the filter.
   NOTE: The volume will depend on the amount of particulate material in the sample: usually between 250 mL and 5 L depending on season and location.
4. When the sample has been measured out, gently wet the filter with filtered seawater. Add the entire sample slowly to the filtration system and rinse the graduated cylinder with filtered seawater to ensure all particles are added to the filter. Rinse the setup with filtered sea water on the sides to ensure all particles are rinsed down onto the filter.
   1. Allow all the seawater to pass through the filter but do not allow the filter to dry out completely as it can disrupt the cells on the filter; break suction as the last of the water disappears.
5. Using clean forceps and a clean pipette, fold the filter in half, then in thirds and then in half lengthwise to roll the filter into a tube. Place it into a clean, labeled 10 mL glass vial.
6. Cover the filter with 2 mL of chloroform. Fill the head space with nitrogen and seal with PTFE tape. Place in a rack in a -20 °C freezer. The samples will be stable at this temperature up to a year.
   NOTE: To relate lipid concentrations to total mass concentrations, a dry weight measurement is also needed. This involves putting a 24 mm pre-weighed filter into a dry weight filtration setup, stirring the sample and taking a small subsample which is filtered onto the small filter. When the filter is nearly dry, about 10 mL of 3.5% ammonium formate is added to the filter. The filter is folded in half, returned to a labeled Petri dish and placed in a freezer.

2. Liquid-liquid extraction of seawater or liquid samples

1. Prepping the sample
   1. Measure a known volume of filtrate into a lipid clean glass graduated cylinder. Place this sample in a clean 1 L separatory funnel. Then add 20 mL of chloroform to the sample and shake for 2 min, venting frequently.
2. Removal of first extract and addition of acid after first separation
   1. Wrap funnel in aluminum foil and wait 5 to 10 min for separation to occur. Peel back the bottom of the foil to see the two layers. Collect the bottom, organic layer through the stopcock into a clean round bottomed flask, being careful not to include any of the top layer. Cap the round bottomed flask under nitrogen and place in the freezer.
   2. Add 0.25 mL of concentrated H₂SO₄ for each liter of sample, to the sample in the separatory funnel and shake the funnel gently.
   3. Add 10 mL of chloroform and shake vigorously for 2 min while venting frequently. Allow the separation to take place.
3. Second and third separations
   1. Wait for the separation and add the bottom layer into the round bottomed flask.
   2. Add a third 10 mL of chloroform, and shake for 2 minutes, again venting frequently. After separation, add the bottom layer to the round bottomed flask.
   3. Transfer the extract in the round bottomed flask to a rotary evaporator, evaporate and transfer into a 2 mL vial.

3. Extraction protocol for solids (modified Folch et al.³ extraction)

1. Setup
   NOTE: The extraction setup requires an insulated container filled with ice. Solvents include methanol, chloroform extracted water and 2:1 chloroform:methanol. All solvents should be placed on ice so that they are cold by the time the extractions are started. The samples also go on ice, so that everything remains cold.
   1. Rinse all tubes and PTFE lined caps 3 times with methanol and then 3 times with chloroform. One centrifuge tube and one 15 mL vial with a cap is needed for each extraction.
   2. Place the sample in a centrifuge tube containing 2 mL of chloroform. The size of the sample depends on the amount of lipid present, approximately 5 to 10 mg of lipid preferred.
2. Grinding and extraction
   1. Add approximately 1 mL of ice-cold methanol and grind the sample into a pulp quickly with a clean homogenizer or PTFE/metal-ended rod.
   2. Wash the rod back into the tube with approximately 1 mL of ice-cold 2:1 chloroform: methanol and then with 1/2 mL of ice-cold chloroform-extracted water. If necessary, use a clean set of forceps to force particles back into the vial before washing. Cap the tube and sonicate the mixture for 4 minutes in an ultrasonic bath (35 - 42 kHz).
3. Double pipetting: Centrifuge the sample for 2 min at 1800 × g. Collect the entire organic layer (bottom layer) using the double pipetting technique in which a 5 ¾" pipette with the bulb on loosely is gently pushed through the two layers, while squeezing the bulb, causing bubbles to come out.
   1. When the bottom of the second layer is reached, use the thumb to remove the bulb without drawing the organic layer into the pipette. Place a 9" pipette inside the 5 ¾" one and remove the bottom layer through the 5 ¾" pipette.
   2. Place the extract in a clean vial. Continue to take off the bottom layer until it is all removed.
4. Washing pipettes: Rinse the 9" pipette into the vial containing the organic layer using 1.5 mL of ice-cold chloroform to wash down the outside of the pipette and 1.5 mL to wash down the inside. Gently turn that pipette while washing and make sure that all the chloroform runs down the pipette into vial.
   1. Rinse the 5 ¾" pipette into the tube containing the aqueous layer in the same way.
5. Sonicate and centrifuge the samples and double pipette when separated, using new pipettes each time. Repeat at least three times and pool all organic layers. After removing the organic layer for the third time, wash both pipettes into the vial containing the organic layer.
6. Using a gentle stream of nitrogen, concentrate down to volume, then seal with PTFE tape and store in a freezer.

4. Developing systems and steps for rod TLC separation of marine lipid classes

1. Prepping the rods for TLC
   1. Blank scan the rods in the automatic FID scanner three times
   2. Spotting a sample: Apply samples and standards with a syringe to the rods at or just below the origin. Dispense 0.5 µL and touch the drop to the rod. Allow to dry before placing the next drop on the same spot. Spot all samples in a line on rods.
   3. Focusing in acetone: Focus samples twice (three times if samples are very concentrated) in 70 mL of acetone. Watch the solvent front as it climbs the rod until the bottom of the spot merges with the top. Remove the rods, dry them for around 5 s, then repeat the procedure to produce a narrow band of lipid material near the bottom of the rod.
   4. Dry and condition the rods in a constant humidity chamber for 5 min. A constant humidity chamber is a desiccator with a saturated solution of calcium chloride under the plate.
2. Sequence leading to the first chromatogram (hydrocarbon to ketone)
   1. First development system: The first development system is hexane:diethyl ether:formic acid, 98.95:1:0.05. Use a syringe to add the formic acid but first rinse the syringe 3× with formic acid. Rinse the formic acid out of the syringe immediately afterwards with chloroform. Use 30 mL of the mixture to wet the paper and rinse the tank. Discard the rinse solution and add the remaining 70 mL to the tank.
      1. Take the racks and gently lower them into the tank. Watch until the solvent front reaches the sample spots, then start the timer. After 25 min, remove the rods from the development chamber, dry in the constant humidity chamber for 5 min, and redevelop in the same solution for another 20 min.
   2. Dry the rods for 5 min in the automatic FID scanner and then scan to the lowest point behind the ketone peak using a PPS scan of 25.
3. Sequence leading to the second chromatogram (triacylglycerol to diacylglycerol):
   1. Condition the rods for 5 min in the constant humidity chamber.
   2. Second development system: The second development system is hexane:diethyl ether:formic acid, 79:20:1. Add ~30 mL to the development tank to wet the paper and rinse the tank. Then discard and develop the rods for 40 min in the remaining 70 mL. For the best separation between the TAG (saturated) and the TAG (polyunsaturated) peaks use a mixture of 79.9:20:0.1, but for separation of the ST and DAG peaks use a mixture of 79:20:1.
   3. Dry and scan to lowest point behind the diacylglycerol peak in a second partial scan using a PPS scan 11.
4. Sequence leading to the third chromatogram (acetone-mobile polar lipid and phospholipid)
   1. Condition the rods for 5 minutes in the constant humidity chamber.
   2. Develop the rods twice for 15 minutes in 70 mL of acetone. Between developments air dry the rods for about 30 seconds.
   3. Condition the rods for 5 minutes in the constant humidity chamber.
   4. Third development system: The third development system is a mixture of chloroform, methanol, and chloroform-extracted water, 50:40:10. Develop the rods twice for 10 minutes in 70 mL of the mixture. Between developments, air dry the rods for about 30 seconds.
   5. Dry and scan entire length of rods.

5. FAME derivatization with H₂SO₄ in MeOH

1. Making the Hilditch reagent
   1. Preparing the methanol: Place 100 mL of MeOH in a clean volumetric flask and then gently sprinkle in anhydrous NaSO₄ until the bottom of the flask is covered. Once covered, invert twice so that any water in the methanol is absorbed by the NaSO₄. After inverting and shaking, let it sit for at least 5 min.
   2. Adding the acid: Slowly decant the methanol into a glass jar (by now the NaSO₄ is a hard lump in the bottom of the flask) and the H₂SO₄ is added. Slowly add 1.5 mL sulphuric acid to the methanol using a pipette. Add a few drops at a time and once all the acid has been added, cap and gently stir to mix. The solution is now ready to be used for derivatives, but it must be made up on a weekly basis.
2. Making the derivatives
   1. Transfer approximately 200 µg of lipids from an extract vial that has had the volume brought up to a known amount, into a clean, 15 mL vial and evaporate under nitrogen to dryness. The amount removed will be determined by the concentration of the sample from TLC/FID. Use a clean pipette to remove the sample.
   2. When the sample has dried, add 1.5 mL of dichloromethane and 3 mL of the newly made Hilditch reagent.
   3. Vortex the sample, and sonicate it in an ultrasonic bath for 4 min to remove lipids that have adhered to the glass vial. Fill the vial with nitrogen, cap, and seal it with PTFE tape and heat at 100°C for 1 hour.
3. Stopping the reaction
   1. Allow the samples to cool completely to room temperature for 10 min after removal from the oven, then open the vials carefully.
   2. Slowly add approximately 0.5 mL saturated sodium bicarbonate solution (9 g/100 mL chloroform-extracted water), then 1.5 mL hexane. Shake or vortex the vial, then let stand so that it separates into 2 layers.
4. Collecting the FAME's
   1. Removing the top layer: Once the derivatization has been halted, and there is clear separation, remove the upper, organic phase and place in a lipid clean 2 mL vial.
   2. Evaporate the solvent in the 2 mL vial to dryness and refill it with hexane to approximately 0.5 mL.
   3. Fill the head space of the vial with nitrogen, cap and seal the vial with PTFE tape, sonicate for another 4 minutes to re-suspend the fatty acids, and then it is ready to go to the GC.
      NOTE: If fatty acid concentrations are required, the aqueous layer must be washed three times with hexane and all the organic layers pooled into the 2 mL vial. This involves adding 2 mL of hexane, vortexing the sample, centrifuging, and removing the organic layer, all repeated 3 times.

Results

As the fastest growing food production sector, aquaculture is evolving in terms of technological innovations and adaptations to meet changing requirements. One of these is to reduce the dependence on wild-sourced fishmeal and fish oil, which provide feed ingredients for many aquaculture species. Terrestrial plant oils are being investigated as sustainable and economical replacements for fish oil in aquafeeds, and the liver is a target tissue for analysis because it is the primary site for lipid metabolism

Discussion

The speed with which the TLC-FID system provides synoptic lipid class information from small samples makes TLC-FID an able tool for screening marine samples before undertaking more involved analytical procedures. Such analyses usually require release of component compounds from lipid extracts and derivatization to increase volatility in the case of gas chromatography. TLC-FID combined with GC-FID has been found to be a powerful combination for extracts of seafood and other foodstuffs¹⁴. For succes...

Materials

Name	Company	Catalog Number	Comments
15 ml vials	VWR	66009-560
1-hexadecanol	Sigma	258741-1G
1-Monopalmitoyl-rac-glycerol	Sigma	M1640-1g
2 ml vials	VWR	46610-722
25 mm glass fibre filters	Fisher	09 874 32A
2ml pipet bulbs	VWR	82024-554
47 mm glass fibre filters	Fisher	09 874 32
5 3/4" pipets	Fisher	1367820A
9" pipets	Fisher	1367820C
Acetone	VWR	CAAX0116-1
Agilent GC-FID 6890	Agilent
Calcium Chloride ANHS 500gm	VWR	CACX0160-1
Caps for 2 ml vials	VWR	46610-712
chloroform	VWR	CACX1054-1
Cholesteryl palmitate	Sigma	C6072-1G
Chromarod S5	Shell USA	3252
Dichloromethane	VWR	CADX0831-1
DL-a-phosphatidylcholine, dipalmotoyl	Sigma	P5911-1g
Ethyl Ether, ACS grade anhydr 4L	VWR	CAEX0190-4
Glyceryl tripalmitate	Sigma	T5888-100MG
Hamilton Syringe 702SNR 25µl	Sigma	58381
Helium	Air Liquide	A0492781
Hexane	VWR	CAHX0296-1
Hydrogen regulator	VWR	55850-484
Iatroscan MK6	Shell USA
Kimwipes	Fisher	066662
Medical Air	Air Liquide	A0464563
Medium nitrile gloves	Fisher	191301597C
Nitrile gloves L	VWR	CA82013-782
Nitrogen	Air Liquide	A0464775
Nitrogen Regulator	VWR	55850-474
Nonadecane	Sigma	74158-1G
Palmitic acid	Sigma	P0500-10G
Repeating dispenser	Sigma	20943
Sodium Bicarbonate 1kg	VWR	CA97062-460
Sodium Sulfate Anhy ACS 500gr	VWR	CA71008-804
Sulfuric acid	VWR	CASX1244-5
Teflon tape	Fisher	14610120
tissue master 125 115V w/7mm homogenator	OMNI International	TM125-115
TLC development tank	Shell USA	3201
UHP hydrogen	Air Liquide	A0492788
VWR solvent repippetter	VWR	82017-766
VWR timer Flashing LED 2 channel	VWR	89140-196
Zebron ZB-Wax GC column	Phenomenex	7HM-G013-11