This research was funded by Natural Sciences and Engineering Research Council of Canada (NSERC) grant number 105379 to C.C. Parrish. Memorial University&#39;s Core Research Equipment &#38; Instrument Training (CREAIT) Network helped fund this publication.

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&#176;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 &#181;m.
<ol>
	<li>Set up the filtration manifold without a filter and rinse the setup with filtered seawater.</li>
	<li>Using clean forceps place a 47 mm glass fiber (GF/C) filter, that has been washed, into the clean filtration system.</li>
	<li>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.</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>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&#160;&#176;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.</li>
</ol>
2. Liquid-liquid extraction of seawater or liquid samples
<ol>
	<li>Prepping the sample
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>Removal of first extract and addition of acid after first separation
	<ol>
		<li>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.</li>
		<li>Add 0.25 mL of concentrated H2SO4 for each liter of sample, to the sample in the separatory funnel and shake the funnel gently.</li>
		<li>Add 10 mL of chloroform and shake vigorously for 2 min while venting frequently. Allow the separation to take place.</li>
	</ol>
	</li>
	<li>Second and third separations
	<ol>
		<li>Wait for the separation and add the bottom layer into the round bottomed flask.</li>
		<li>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.</li>
		<li>Transfer the extract in the round bottomed flask to a rotary evaporator, evaporate and transfer into a 2 mL vial.</li>
	</ol>
	</li>
</ol>
3. Extraction protocol for solids (modified Folch et al.3 extraction)
<ol>
	<li>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.
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Grinding and extraction
	<ol>
		<li>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.</li>
		<li>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).</li>
	</ol>
	</li>
	<li>Double pipetting: Centrifuge the sample for 2 min at 1800 &#215; g. Collect the entire organic layer (bottom layer) using the double pipetting technique in which a 5 &#190;&#34; pipette with the bulb on loosely is gently pushed through the two layers, while squeezing the bulb, causing bubbles to come out.
	<ol>
		<li>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&#34; pipette inside the 5 &#190;&#34; one and remove the bottom layer through the 5 &#190;&#34; pipette.</li>
		<li>Place the extract in a clean vial. Continue to take off the bottom layer until it is all removed.</li>
	</ol>
	</li>
	<li>Washing pipettes: Rinse the 9&#34; 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.
	<ol>
		<li>Rinse the 5 &#190;&#34; pipette into the tube containing the aqueous layer in the same way.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Using a gentle stream of nitrogen, concentrate down to volume, then seal with PTFE tape and store in a freezer.</li>
</ol>
4. Developing systems and steps for rod TLC separation of marine lipid classes
<ol>
	<li>Prepping the rods for TLC
	<ol>
		<li>Blank scan the rods in the automatic FID scanner three times</li>
		<li>Spotting a sample: Apply samples and standards with a syringe to the rods at or just below the origin. Dispense 0.5 &#181;L 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.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Sequence leading to the first chromatogram (hydrocarbon to ketone)
	<ol>
		<li>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&#215; with formic acid. Rinse the formic acid out&#160;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.
		<ol>
			<li>Take the racks and gently lower them into the tank. Watch until the solvent front reaches the sample&#160;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.</li>
		</ol>
		</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Sequence leading to the second chromatogram (triacylglycerol to diacylglycerol):
	<ol>
		<li>Condition the rods for 5 min in the constant humidity chamber.</li>
		<li>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.</li>
		<li>Dry and scan to lowest point behind the diacylglycerol peak in a second partial scan using a PPS scan 11.</li>
	</ol>
	</li>
	<li>Sequence leading to the third chromatogram (acetone-mobile polar lipid and phospholipid)
	<ol>
		<li>Condition the rods for 5 minutes in the constant humidity chamber.</li>
		<li>Develop the rods twice for 15 minutes in 70 mL of acetone. Between developments air dry the rods for about 30 seconds.</li>
		<li>Condition the rods for 5 minutes in the constant humidity chamber.</li>
		<li>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.</li>
		<li>Dry and scan entire length of rods.</li>
	</ol>
	</li>
</ol>
5. FAME derivatization with H2SO4 in MeOH
<ol>
	<li>Making the Hilditch reagent
	<ol>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Making the derivatives
	<ol>
		<li>Transfer approximately 200 &#181;g 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.</li>
		<li>When the sample has dried, add 1.5 mL of dichloromethane and 3 mL of the newly made Hilditch reagent.</li>
		<li>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&#176;C for 1 hour.</li>
	</ol>
	</li>
	<li>Stopping the reaction
	<ol>
		<li>Allow the samples to cool completely to room temperature for 10 min after removal from the oven, then open the vials carefully.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Collecting the FAME&#39;s
	<ol>
		<li>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.</li>
		<li>Evaporate the solvent in the 2 mL vial to dryness and refill it with hexane to approximately 0.5 mL.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>

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</ol>

The authors have no competing financial interests.

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...

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 &#39;dissolved&#39; and &#39;particulate&#39; 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 esteri&#64257;ed 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).

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

determination of total lipid and lipid classes in marine samples

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 &#39;dissolved&#39; and &#39;particulate&#39; 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).

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<sup class="xref...

Watch this Scientific Journal Video about Determination of Total Lipid and Lipid Classes in Marine Samples at JoVE.com

Determination of Total Lipid and Lipid Classes in Marine Samples

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.

Lipids are largely composed of carbon and hydrogen and, therefore, provide a greater specific energy than other organic macromolecules in the sea. Being ...

determination-of-total-lipid-and-lipid-classes-in-marine-samples

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4.5K Views.  Memorial University of Newfoundland. 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. 

Video: Determination of Total Lipid and Lipid Classes in Marine Samples

Analysis of Fatty Acid Content and Composition in Microalgae

A method to determine the content and composition of total fatty acids present in microalgae is described. Fatty acids are a major constituent of microalgal biomass. These fatty acids can be present in different acyl-lipid classes. Especially the fatty acids present in triacylglycerol (TAG) are of commercial interest, because they can be used for production of transportation fuels, bulk chemicals, nutraceuticals (&omega;-3 fatty acids), and food commodities. To develop commercial applications, reliable analytical methods for quantification of fatty acid content and composition are needed. Microalgae are single cells surrounded by a rigid cell wall. A fatty acid analysis method should provide sufficient cell disruption to liberate all acyl lipids and the extraction procedure used should be able to extract all acyl lipid classes.
With the method presented here all fatty acids present in microalgae can be accurately and reproducibly identified and quantified using small amounts of sample (5 mg) independent of their chain length, degree of unsaturation, or the lipid class they are part of.
This method does not provide information about the relative abundance of different lipid classes, but can be extended to separate lipid classes from each other.
The method is based on a sequence of mechanical cell disruption, solvent based lipid extraction, transesterification of fatty acids to fatty acid methyl esters (FAMEs), and quantification and identification of FAMEs using gas chromatography (GC-FID). A TAG internal standard (tripentadecanoin) is added prior to the analytical procedure to correct for losses during extraction and incomplete transesterification.

A method for the determination of fatty acid content and composition in microalgae based on mechanical cell disruption, solvent based lipid extraction, transesterification, and quantification and identification of fatty acids using gas chromatography is described. A tripentadecanoin internal standard is used to compensate for the possible losses during extraction and incomplete transesterification.

A method to determine the content and composition of total fatty acids present in microalgae is described. Fatty acids are a major constituent of ...

A Lipid Extraction and Analysis Method for Characterizing Soil Microbes in Experiments with Many Samples

Microbial communities are important drivers and regulators of ecosystem processes. To understand how management of ecosystems may affect microbial communities, a relatively precise but effort-intensive technique to assay microbial community composition is phospholipid fatty acid (PLFA) analysis. PLFA was developed to analyze phospholipid biomarkers, which can be used as indicators of microbial biomass and the composition of broad functional groups of fungi and bacteria. It has commonly been used to compare soils under alternative plant communities, ecology, and management regimes. The PLFA method has been shown to be sensitive to detecting shifts in microbial community composition.
An alternative method, fatty acid methyl ester extraction and analysis (MIDI-FA) was developed for rapid extraction of total lipids, without separation of the phospholipid fraction, from pure cultures as a microbial identification technique. This method is rapid but is less suited for soil samples because it lacks an initial step separating soil particles and begins instead with a saponification reaction that likely produces artifacts from the background organic matter in the soil. 
This article describes a method that increases throughput while balancing effort and accuracy for extraction of lipids from the cell membranes of microorganisms for use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure in studies with many samples. The method combines the accuracy achieved through PLFA profiling by extracting and concentrating soil lipids as a first step, and a reduction in effort by saponifying the organic material extracted and processing with the MIDI-FA method as a second step.

The article describes a method that increases throughput while balancing effort and accuracy for extraction of lipids from the cell membranes of microorganisms for use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure in studies with many samples.

Microbial communities are important drivers and regulators of ecosystem processes. To understand how management of ecosystems may affect microbial ...

Fatty Acid 13C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

Fatty acids (FAs) are useful biomarkers in food web ecology because they are typically assimilated as a complete molecule and transferred into consumer tissue with minor or no modification, allowing the dietary routing between different trophic levels. However, the FA trophic marker approach is still hampered by the limited knowledge in lipid metabolism of the soil fauna. This study used entirely labelled palmitic acid (13C16:0, 99 atom%) as a tracer in fatty acid metabolism pathways of two widespread soil Collembola, Protaphorura fimata and Heteromurus nitidus. In order to investigate the fate and metabolic modifications of this precursor, a method of isotopologue profiling is presented, performed by mass spectrometry using single ion monitoring. Moreover, the upstream laboratory feeding experiment is described, as well as the extraction and methylation of dominant lipid fractions (neutral lipids, phospholipids) and the related formula and calculations. Isotopologue profiling does not only yield the overall 13C enrichment in fatty acids derived from the 13C labeled precursor but also produces the pattern of isotopologues exceeding the mass of the parent ion (i.e., the FA molecular ion M+) of each labeled FA by one or more mass units (M+1, M+2, M+3, etc.). This knowledge allows conclusions on the ratio of dietary routing of an entirely consumed FA in comparison to de novo biosynthesis. The isotopologue profiling is suggested as a useful tool for evaluation of fatty acid metabolism in soil animals to disentangle trophic interactions.

Fatty Acid 13C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

The fatty acid trophic marker approach, i.e., the assimilation of fatty acids as entire molecule and transfer into consumer tissue with no or minor modification, is hampered by knowledge gaps in fatty acid metabolism of small soil invertebrates. Isotopologue profiling is provided as a valuable tool to disentangle trophic interactions.

Fatty acids (FAs) are useful biomarkers in food web ecology because they are typically assimilated as a complete molecule and transferred into consumer ...

Biochemistry

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of biological membranes, energy storage and production, cellular signaling, vesicular transport of proteins, organelle biogenesis, and regulated cell death. Because the budding yeast Saccharomyces cerevisiae is a unicellular eukaryote amenable to thorough molecular analyses, its use as a model organism helped uncover mechanisms linking lipid metabolism and intracellular transport to complex biological processes within eukaryotic cells. The availability of a versatile analytical method for the robust, sensitive, and accurate quantitative assessment of major classes of lipids within a yeast cell is crucial for getting deep insights into these mechanisms. Here we present a protocol to use liquid chromatography coupled with tandem mass spectrometry (LC-MS/MS) for the quantitative analysis of major cellular lipids of S. cerevisiae. The LC-MS/MS method described is versatile and robust. It enables the identification and quantification of numerous species (including different isobaric or isomeric forms) within each of the 10 lipid classes. This method is sensitive and allows identification and quantitation of some lipid species at concentrations as low as 0.2 pmol/&#181;L. The method has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. The use of alternative mobile phase additives for electrospray ionization mass spectrometry in this method can increase the efficiency of ionization for some lipid species and can be therefore used to improve their identification and quantitation.

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

We present a protocol using liquid chromatography coupled with tandem mass spectrometry to identify and quantify major cellular lipids in Saccharomyces cerevisiae. The described method for a quantitative assessment of major lipid classes within a yeast cell is versatile, robust, and sensitive.

Lipids are structurally diverse amphipathic molecules that are insoluble in water. Lipids are essential contributors to the organization and function of ...

A Quantitative Assessment of The Yeast Lipidome using Electrospray Ionization Mass Spectrometry

Lipids are one of the major classes of biomolecules and play important roles membrane dynamics, energy storage, and signalling1-4. The budding yeast Saccharomyces cerevisiae, a genetically and biochemically manipulable unicellular eukaryote with annotated genome and very simple lipidome, is a valuable model for studying biological functions of various lipid species in multicellular eukaryotes2,3,5. S. cerevisiae has 10 major classes of lipids with chain lengths mainly of 16 or 18 carbon atoms and either zero or one degree of unsaturation6,7. Existing methods for lipid identification and quantification - such as high performance liquid chromatography, thin-layer chromatography, fluorescence microscopy, and gas chromatography followed by MS - are well established but have low sensitivity, insufficiently separate various molecular forms of lipids, require lipid derivitization prior to analysis, or can be quite time consuming. Here we present a detailed description of our experimental approach to solve these inherent limitations by using survey-scan ESI/MS for the identification and quantification of the entire complement of lipids in yeast cells. The described method does not require chromatographic separation of complex lipid mixtures recovered from yeast cells, thereby greatly accelerating the process of data acquisition. This method enables lipid identification and quantification at the concentrations as low as g/ml and has been successfully applied to assessing lipidomes of whole yeast cells and their purified organelles. Lipids extraction from whole yeast cells for using this method of lipid analysis takes two to three hours. It takes only five to ten minutes to run each sample of extracted and dried lipids on a Q-TOF mass spectrometer equipped with a nano-electrospray source.

We describe a new quantitative lipidomics method for identifying numerous lipid species in yeast using survey-scan electrospray ionization mass spectrometry (ESI/MS). This method exceeds currently available methods for lipid identification and quantification in the ability to resolve various molecular forms of lipids, sensitivity, and speed.

Lipids are one of the major classes of biomolecules and play important roles membrane dynamics, energy storage, and signalling1-4. The budding yeast ...

Biology

Shotgun Lipidomics of Rodent Tissues

Lipids play a vital role as essential components of all prokaryotic and eukaryotic cells. Constant technological improvements in mass spectrometry have made lipidomics a powerful analytical tool for monitoring tissue lipidome compositions in homeostatic as well as disease states. This paper presents a step-by-step protocol for a shotgun lipid analysis method that supports the simultaneous detection and quantification of a few hundred molecular lipid species in different tissue and biofluid samples at high throughput. This method leverages automated nano-flow direct injection of a total lipid extract spiked with labeled internal standards without chromatographic separation into a high-resolution mass spectrometry instrument. Starting from sub-microgram amounts of rodent tissue, the MS analysis takes 10 min per sample and covers up to 400 lipids from 14 lipid classes in mouse lung tissue. The method presented here is well suited for studying disease mechanisms and identifying and quantifying biomarkers that indicate early toxicity or beneficial effects within rodent tissues.

Shotgun mass spectrometry-based lipidomics delivers a sensitive quantitative snapshot of a broad spectrum of lipid classes simultaneously in a single measurement from various rodent tissues.

Lipids play a vital role as essential components of all prokaryotic and eukaryotic cells. Constant technological improvements in mass spectrometry have ...

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

The nematode C. elegans has emerged as an important model for the study of conserved genetic pathways regulating fat metabolism as it relates to human obesity and its associated pathologies. Several previous methodologies developed for the visualization of C. elegans triglyceride-rich fat stores have proven to be erroneous, highlighting cellular compartments other than lipid droplets. Other methods require specialized equipment, are time-consuming, or yield inconsistent results. We introduce a rapid, reproducible, fixative-based Nile red staining method for the accurate and rapid detection of neutral lipid droplets in C. elegans. A short fixation step in 40% isopropanol makes animals completely permeable to Nile red, which is then used to stain animals. Spectral properties of this lipophilic dye allow it to strongly and selectively fluoresce in the yellow-green spectrum only when in a lipid-rich environment, but not in more polar environments. Thus, lipid droplets can be visualized on a fluorescent microscope equipped with simple GFP imaging capability after only a brief Nile red staining step in isopropanol. The speed, affordability, and reproducibility of this protocol make it ideally suited for high throughput screens. We also demonstrate a paired method for the biochemical determination of triglycerides and phospholipids using gas chromatography mass-spectrometry. This more rigorous protocol should be used as confirmation of results obtained from the Nile red microscopic lipid determination. We anticipate that these techniques will become new standards in the field of C. elegans metabolic research.

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

We present robust biochemical and microscopic methods for studying Caenorhabditis elegans lipid stores. A rapid, simple, fixing-staining procedure for fluorescent lipid droplet imaging leverages the spectral properties of the lipophilic dye Nile red. We then present biochemical measurement of triglycerides and phospholipids using solid phase extraction and gas chromatography-mass spectrometry.

The nematode C. elegans has emerged as an important  model for the study of conserved genetic pathways regulating fat metabolism as  it relates to human ...

Quantitative Determination of De Novo Fatty Acid Synthesis in Brown Adipose Tissue Using Deuterium Oxide

Fatty acid synthesis is a complex and highly energy demanding metabolic pathway with important functional roles in the control of whole-body metabolic homeostasis and other physiological and pathological processes. Contrary to other key metabolic pathways, such as glucose disposal, fatty acid synthesis is not routinely functionally assessed, leading to incomplete interpretations of metabolic status. In addition, there is a lack of publicly available detailed protocols suitable for newcomers in the field. Here, we describe an inexpensive quantitative method utilizing deuterium oxide and gas chromatography mass spectrometry (GCMS) for the analysis of total fatty acid de novo synthesis in brown adipose tissue in vivo. This method measures the synthesis of the products of fatty acid synthase independently of a carbon source, and it is potentially useful for virtually any tissue, in any mouse model, and under any external perturbation. Details on the sample preparation for GCMS and downstream calculations are provided. We focus on the analysis of brown fat due to its high levels of de novo fatty acid synthesis and critical roles in maintaining metabolic homeostasis.

Quantitative Determination of De Novo Fatty Acid Synthesis in Brown Adipose Tissue Using Deuterium Oxide

Here, we present an inexpensive quantitative method utilizing deuterium oxide and gas chromatography mass spectrometry (GCMS) for the analysis of total fatty acid de novo lipogenesis in brown adipose tissue in vivo.

Fatty acid synthesis is a complex and highly energy demanding metabolic pathway with important functional roles in the control of whole-body metabolic ...

Sonication Extraction of Lipid Biomarkers from Sediment

Source: Laboratory of Jeff Salacup&#160;- University of Massachusetts Amherst
The material comprising the living &#34;organic&#34; share of any ecosystem (leaves, fungi, bark, tissue; Figure 1) differs fundamentally from the material of the non-living &#34;inorganic&#34; share (rocks and their constituent minerals, oxygen, water, metals). Organic material contains carbon linked to a series of other carbon and hydrogen molecules (Figure 2), which distinguishes it from inorganic material. Carbon&#39;s wide valency range (-4 to +4) allows it to form up to four separate covalent bonds with neighboring atoms, usually C, H, O, N, S, and P. It can also share up to three covalent bonds with a single other atom, such as the triple bond in the often-poisonous cyanide, or nitrile, group. Over the past 4.6 billion years, this flexibility has led to an amazing array of chemical structures, which vary in size, complexity, polarity, shape, and function. The scientific field of organic geochemistry is concerned with the identification and characterization of the whole range of detectable organic compounds, called biomarkers, produced by life on this planet, as well as others, through geologic time.
<img alt="Figure 1" src="/files/ftp_upload/10055/10055fig1.jpg" /> 
Figure 1. Organic material, such as trees, leaves, and moss, are chemically and visually distinct from inorganic material, such as pavement.

Source: Laboratory of Jeff Salacup&#160;- University of Massachusetts Amherst
The material comprising the living &#34;organic&#34; share of any ecosystem ...

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Soxhlet Extraction of Lipid Biomarkers from Sediment

Source: Laboratory of Jeff Salacup&#160;- University of Massachusetts Amherst
Every lab needs standards that track the performance, accuracy, and precision of its instruments over time to ensure a measurement made today is the same as a measurement made a year from now (Figure 1). Because standards must test the performance of instruments over a long period of time, large volumes of the standards are often required. Many chemical standards can be purchased from retail scientific companies, like Sigma-Aldrich and Fisher. However, some compounds that occur in nature and that are relevant to paleoclimatic studies have not yet been isolated and purified for purchase. Therefore, these compounds need to be extracted from natural samples, and because of the large volumes of standards required, large volumes of sediment need to be extracted. The Accelerated Solvent Extraction (Dionex) and sonication extractions are not appropriate for the extraction of such large sediment volumes. In these circumstances, a Soxhlet extraction is used.
<img alt="Figure 1" src="/files/ftp_upload/10096/10096fig1.jpg" /> 
Figure 1. Schematic depicting how chemical standard tracks the performance of an instrument through time. The dashed line represents a 1:1 relationship between the accepted and measured (on the instrument) value of a variable. Each star is a weekly measurement of the chemical standard. Green stars represent standards that are accurate. Red stars reflect those that are not accurate indicating that the instrument requires corrective maintenance.

Determination of Total Lipid and Lipid Classes in Marine Samples

Summary

Explore More Videos

Analysis of Fatty Acid Content and Composition in Microalgae

A Lipid Extraction and Analysis Method for Characterizing Soil Microbes in Experiments with Many Samples

Fatty Acid ¹³C Isotopologue Profiling Provides Insight into Trophic Carbon Transfer and Lipid Metabolism of Invertebrate Consumers

Quantitative Analysis of the Cellular Lipidome of Saccharomyces Cerevisiae Using Liquid Chromatography Coupled with Tandem Mass Spectrometry

A Quantitative Assessment of The Yeast Lipidome using Electrospray Ionization Mass Spectrometry

Shotgun Lipidomics of Rodent Tissues

Biochemical and High Throughput Microscopic Assessment of Fat Mass in Caenorhabditis Elegans

Quantitative Determination of De Novo Fatty Acid Synthesis in Brown Adipose Tissue Using Deuterium Oxide

Sonication Extraction of Lipid Biomarkers from Sediment

Soxhlet Extraction of Lipid Biomarkers from Sediment