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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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).

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			<td>chloroform</td>
			<td>VWR</td>
			<td>CACX1054-1</td>
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			<td>Cholesteryl palmitate</td>
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			<td>DL-a-phosphatidylcholine, dipalmotoyl</td>
			<td>Sigma</td>
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			<td>Sigma</td>
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			<td>Phenomenex</td>
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</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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4.3K 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

A Noninvasive Method For In situ Determination of Mating Success in Female American Lobsters (Homarus americanus)

Despite being one of the most productive fisheries in the Northwest Atlantic, much remains unknown about the natural reproductive dynamics of American lobsters. Recent work in exploited crustacean populations (crabs and lobsters) suggests that there are circumstances where mature females are unable to achieve their full reproductive potential due to sperm limitation. To examine this possibility in different regions of the American lobster fishery, a reliable and noninvasive method was developed for sampling large numbers of female lobsters at sea. This method involves inserting a blunt-tipped needle into the female&#39;s seminal receptacle to determine the presence or absence of a sperm plug and to withdraw a sample that can be examined for the presence of sperm. A series of control studies were conducted at the dock and in the laboratory to test the reliability of this technique. These efforts entailed sampling 294 female lobsters to confirm that the presence of a sperm plug was a reliable indicator of sperm within the receptacle and thus, mating. This paper details the methodology and the results obtained from a subset of the total females sampled. Of the 230 female lobsters sampled from George&#39;s Bank and Cape Ann, MA (size range = 71-145 mm in carapace length), 90.3% were positive for sperm. Potential explanations for the absence of sperm in some females include: immaturity (lack of physiological maturity), breakdown of the sperm plug after being used to fertilize a clutch of eggs, and lack of mating activity. The surveys indicate that this technique for examining the mating success of female lobsters is a reliable proxy that can be used in the field to document reproductive activity in natural populations.

A Noninvasive Method For In situ Determination of Mating Success in Female American Lobsters Homarus americanus

Fishery-induced changes to exploited crustacean fisheries, such as the American lobster fishery, could potentially influence their reproductive dynamics, leading to a reduction in mating success. This study&#39;s goal was to develop a noninvasive method for ascertaining the mating success of female lobsters that may be physiologically or functionally mature.

Despite being one of the most productive fisheries in the Northwest Atlantic, much remains unknown about the natural reproductive dynamics of American ...

A Fish-feeding Laboratory Bioassay to Assess the Antipredatory Activity of Secondary Metabolites from the Tissues of Marine Organisms

Marine chemical ecology is a young discipline, having emerged from the collaboration of natural products chemists and marine ecologists in the 1980s with the goal of examining the ecological functions of secondary metabolites from the tissues of marine organisms. The result has been a progression of protocols that have increasingly refined the ecological relevance of the experimental approach. Here we present the most up-to-date version of a fish-feeding laboratory bioassay that enables investigators to assess the antipredatory activity of secondary metabolites from the tissues of marine organisms. Organic metabolites of all polarities are exhaustively extracted from the tissue of the target organism and reconstituted at natural concentrations in a nutritionally appropriate food matrix. Experimental food pellets are presented to a generalist predator in laboratory feeding assays to assess the antipredatory activity of the extract. The procedure described herein uses the bluehead, Thalassoma bifasciatum, to test the palatability of Caribbean marine invertebrates; however, the design may be readily adapted to other systems. Results obtained using this laboratory assay are an important prelude to field experiments that rely on the feeding responses of a full complement of potential predators. Additionally, this bioassay can be used to direct the isolation of feeding-deterrent metabolites through bioassay-guided fractionation. This feeding bioassay has advanced our understanding of the factors that control the distribution and abundance of marine invertebrates on Caribbean coral reefs and may inform investigations in diverse fields of inquiry, including pharmacology, biotechnology, and evolutionary ecology.

This bioassay employs a model predatory fish to assess the presence of feeding-deterrent metabolites from organic extracts of the tissues of marine organisms at natural concentrations using a nutritionally comparable food matrix.

Marine chemical ecology is a young discipline, having emerged from the collaboration of natural products chemists and marine ecologists in the 1980s with ...

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and detritus, and excreting particulate and dissolved compounds, they serve as important agents for benthic-pelagic coupling. Accurately measuring the compounds removed and excreted by suspension feeders (such as sponges, ascidians, polychaetes, bivalves) is crucial for the study of their physiology, metabolism, and feeding ecology, and is fundamental to determine the ecological relevance of the nutrient fluxes mediated by these organisms. However, the assessment of the rate by which suspension feeders process particulate and dissolved compounds in nature is restricted by the limitations of the currently available methodologies. Our goal was to develop a simple, reliable, and non-intrusive method that would allow clean and controlled water sampling from a specific point, such as the excurrent aperture of benthic suspension feeders, in situ. Our method allows simultaneous sampling of inhaled and exhaled water of the studied organism by using minute tubes installed on a custom-built manipulator device and carefully positioned inside the exhalant orifice of the sampled organism. Piercing a septum on the collecting vessel with a syringe needle attached to the distal end of each tube allows the external pressure to slowly force the sampled water into the vessel through the sampling tube. The slow and controlled sampling rate allows integrating the inherent patchiness in the water while ensuring contamination free sampling. We provide recommendations for the most suitable filtering devices, collection vessel, and storing procedures for the analyses of different particulate and dissolved compounds. The VacuSIP system offers a reliable method for the quantification of undisturbed suspension feeder metabolism in natural conditions that is cheap and easy to learn and apply to assess the physiology and functional role of filter feeders in different ecosystems.

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

We introduce the VacuSIP, a simple, non-intrusive, and reliable method for clean and accurate point sampling of water. The system was developed and evaluated for the simultaneous collection of the water inhaled and exhaled by benthic suspension feeders in situ, to cleanly measure removal and excretion of particulate and dissolved compounds.

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and ...

Ecotoxicological Method with Marine Bacteria Vibrio anguillarum to Evaluate the Acute Toxicity of Environmental Contaminants

Bacteria are an important component of the ecosystem, and microbial community alterations can have a significant effect on biogeochemical cycling and food webs. Toxicity testing based on microorganisms are widely used because they are relatively quick, reproducible, cheap, and are not associated with ethical issues. Here, we describe an ecotoxicological method to evaluate the biological response of the marine bacterium Vibrio anguillarum. This method assesses the acute toxicity of chemical compounds, including new contaminants such as nanoparticles, as well as environmental samples. The endpoint is the reduction of bacterial culturability (i.e., the capability to replicate and form colonies) due to exposure to a toxicant. This reduction can be generally referred to as mortality. The test allows for the determination of the LC50, the concentration that causes a 50% decrease of bacteria actively replicating and forming colonies, after a 6 h exposure. The culturable bacteria are counted in terms of colony forming units (CFU), and the &#34;mortality&#34; is evaluated and compared to the control. In this work, the toxicity of copper sulphate (CuSO4) was evaluated. A clear dose-response relationship was observed, with a mean LC50 of 1.13 mg/L, after three independent tests. This protocol, compared to existing methods with microorganisms, is applicable in a wider range of salinity and has no limitations for colored/turbid samples. It uses saline solution as the exposure medium, avoiding any possible interferences of growth medium with the investigated contaminants. The LC50 calculation facilitates comparisons with other bioassays commonly applied to ecotoxicological assessments of the marine environment.

Ecotoxicological Method with Marine Bacteria Vibrio anguillarum to Evaluate the Acute Toxicity of Environmental Contaminants

This work describes a new protocol to assess the ecotoxicity of pollutants, including emerging contaminants such as nanomaterials, using the marine bacterium Vibrio anguillarum. This method allows for the determination of the LC50, or mortality, the concentration that causes a 50% decrease in bacteria culturability, after a 6 h exposure.

Bacteria are an important component of the ecosystem, and microbial community alterations can have a significant effect on biogeochemical cycling and food ...

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 ...

Preparation of Authigenic Pyrite from Methane-bearing Sediments for In Situ Sulfur Isotope Analysis Using SIMS

Different sulfur isotope compositions of authigenic pyrite typically result from the sulfate-driven anaerobic oxidation of methane (SO4-AOM) and organiclastic sulfate reduction (OSR) in marine sediments. However, unravelling the complex pyritization sequence is a challenge because of the coexistence of different sequentially formed pyrite phases. This manuscript describes a sample preparation procedure that enables the use of secondary ion mass spectroscopy (SIMS) to obtain in situ &#948;34S values of various pyrite generations. This allows researchers to constrain how SO4-AOM affects pyritization in methane-bearing sediments. SIMS analysis revealed an extreme range in &#948;34S values, spanning from -41.6 to +114.8&#8240;, which is much wider than the range of &#948;34S values obtained by the traditional bulk sulfur isotope analysis of the same samples. Pyrite in the shallow sediment mainly consists of 34S-depleted framboids, suggesting early diagenetic formation by OSR. Deeper in the sediment, more pyrite occurs as overgrowths and euhedral crystals, which display much higher SIMS &#948;34S values than the framboids. Such 34S-enriched pyrite is related to enhanced SO4-AOM at the sulfate-methane transition zone, postdating OSR. High-resolution in situ SIMS sulfur isotope analyses allow for the reconstruction of the pyritization processes, which cannot be resolved by bulk sulfur isotope analysis.

Preparation of Authigenic Pyrite from Methane-bearing Sediments for In Situ Sulfur Isotope Analysis Using SIMS

Analyses of the sulfur isotopic composition (&#948;34S) of pyrite from methane-bearing sediments have typically focused on bulk samples. Here, we applied secondary ion mass spectroscopy to analyze the &#948;34S values of various pyrite generations to understand the diagenetic history of pyritization.

Different sulfur isotope compositions of authigenic pyrite typically result from the sulfate-driven anaerobic oxidation of methane (SO4-AOM) and ...

An Ultra-clean Multilayer Apparatus for Collecting Size Fractionated Marine Plankton and Suspended Particles

The distributions of many trace elements in the ocean are strongly associated with the growth, death, and re-mineralization of marine plankton and those of suspended/sinking particles. Here, we present an all plastic (Polypropylene and Polycarbonate), multi-layer filtration system for collection of suspended particulate matter (SPM) at sea. This ultra-clean sampling device has been designed and developed specifically for trace element studies. Meticulous selection of all non-metallic materials and utilization of an in-line flow-through procedure minimizes any possible metal contamination during sampling. This system has been successfully tested and tweaked for determining trace metals (e.g., Fe, Al, Mn, Cd, Cu, Ni) on particles of varying size in coastal and open ocean waters. Results from the South China Sea at the South East Asia Time-Series (SEATS) station indicate that diurnal variations and spatial distribution of plankton in the euphotic zone can be easily resolved and recognized. Chemical analysis of size-fractionated particles in surface waters of the Taiwan Strait suggests that the larger particles (&#62;153 &#181;m) were mostly biologically derived, while the smaller particles (10 - 63 &#181;m) were mostly composed of inorganic matter. Apart from Cd, the concentrations of metals (Fe, Al, Mn, Cu, Ni) decreased with increasing size.

Plankton and suspended particles play a major role in the biogeochemical cycles in the ocean. Here, we provide an ultra-clean, low stress method for the collection of various sizes of particles and plankton at sea with the capability of handling large volumes of seawater.

The distributions of many trace elements in the ocean are strongly associated with the growth, death, and re-mineralization of marine plankton and those ...

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly relevant as a model for the studies of osmoregulatory mechanisms because of its extreme tolerance to low salinity. It is also widely used as a model of settling biology, in particular in relation to antifouling research. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae for studies. A protocol for the all-year-round culturing of B. improvisus has been developed and a detailed description of all steps in the production line is outlined (i.e., the establishment of adult cultures on panels, the collection and rearing of barnacle larvae, and the administration of feed for adults and larvae). The description also provides guidance on troubleshooting and discusses critical parameters (e.g., the removal of contamination, the production of high-quality feed, the manpower needed, and the importance of high-quality seawater). Each batch from the culturing system maximally yields roughly 12,000 nauplii and can deliver four batches in a week, so up to almost 50,000 larvae per week can be produced. The method used to culture B. improvisus is, probably, to a large extent also applicable to other marine invertebrates with free-swimminglarvae. Protocols are presented for the dissection of various tissues from adults as well as the production of high-quality RNA for studies on gene expression. It is also described how cultured adults and reared cyprids can be utilized in a wide array of experimental designs for examining gene expression in relation to external factors. The use of cultured barnacles in gene expression is illustrated with studies of possible osmoregulatory roles of Na+/K+ ATPase and aquaporins.

The Barnacle Balanus improvisus as a Marine Model - Culturing and Gene Expression

The barnacle Balanus (Amphibalanus) improvisus is a model for studying osmoregulation and antifouling. However, natural seasonal spawning yields an unpredictable supply of cyprid larvae. Here, a protocol for the all-year-round culturing of B. improvisus is described, including the production of larvae. The use of cultured barnacles in gene expression studies is illustrated.

Barnacles are marine crustaceans with a sessile adult and free-swimming, planktonic larvae. The barnacle Balanus (Amphibalanus) improvisus is particularly ...

In Situ Hybridization Techniques for Paraffin-Embedded Adult Coral Samples

Corals are important ocean invertebrates that are critical for overall ocean health as well as human health. However, due to human impacts such as rising ocean temperatures and ocean acidification, corals are increasingly under threat. To tackle these challenges, advances in cell and molecular biology have proven to be crucial for diagnosing the health of corals. Modifying some of the techniques commonly used in human medicine could greatly improve researchers&#39; ability to treat and save corals. To address this, a protocol for in situ hybridization used primarily in human medicine and evolutionary developmental biology has been adapted for use in adult corals under stress.
The purpose of this method is to visualize the spatial expression of an RNA probe in adult coral tissue that has been embedded in paraffin and sectioned onto glass slides. This method focuses on removal of the paraffin and rehydration of the sample, pretreatment of the sample to ensure permeability of the sample, pre-hybridization incubation, hybridization of the RNA probe, and visualization of the RNA probe. This is a powerful method when using non-model organisms to discover where specific genes are expressed, and the protocol can be easily adapted for other non-model organisms. However, the method is limited in that it is primarily qualitative, because expression intensity can vary depending on the amount of time used during the visualization step and the concentration of the probe. Furthermore, patience is required, as this protocol can take up to 5 days (and in many cases, longer) depending on the probe being used. Finally, non-specific background staining is common, but this limitation can be overcome.

In Situ Hybridization Techniques for Paraffin-Embedded Adult Coral Samples

The goal of this protocol is to perform in situ hybridization on adult coral samples that have been embedded in paraffin and sectioned onto glass slides. This is a qualitative method used to visualize the spatial expression of an RNA anti-sense probe in paraffin-embedded tissues.

Corals are important ocean invertebrates that are critical for overall ocean health as well as human health. However, due to human impacts such as rising ...

Individual Culturing of Tigriopus Copepods and Quantitative Analysis of Their Mate-guarding Behavior

Copepods of the genus Tigriopus, which are common zooplankton in rocky tide pools, show precopulatory mate-guarding behavior where a male clasps a potential mate to form a pair. While this phenomenon has attracted interest of researchers, methods for its analysis have not been well described. Here we describe procedures for: 1) individual culturing and staging of Tigriopus juveniles and adults, and 2) video-based analysis of their mate-guarding behavior. The culturing method enables experimental control of paring experience of animals as well as the ability to track their development before behavioral tests. The analysis method allows quantitative evaluation of several aspects of the mate-guarding behavior, including capturing attempts by males and swimming trajectory of mate-guarding pairs. Although these methods were originally established for ethological studies on Tigriopus, with proper modifications they can also be applied to studies of other zooplankton in different research fields, such as physiology, toxicology, and ecological genetics.

Individual Culturing of Tigriopus Copepods and Quantitative Analysis of Their Mate-guarding Behavior

Mate-guarding behavior plays an important role in reproduction of intertidal copepods of the genus Tigriopus. However, methods for studying this behavior have not been well described. Here we describe methods for: 1) individual culture of virgin Tigriopus animals, and 2) quantitative analysis of their mate-guarding behavior.