We are grateful to the many persons who have contributed accumulated knowledge to this project over the years including G.-A. Paffenh&#246;fer and D. Deibel who originally developed these protocols. M. K&#246;ster, and L. Lamboley have also contributed significantly to the development of these procedures.&#160; N.B. L&#243;pez-Figueroa and &#193;.E. Rodr&#237;guez-Santiago generated the estimates of doliolid abundance provided in Table 1. This study was supported in part by the US National Science Foundation awards OCE 082599, 1031263 to MEF, collaborative projects OCE 1459293 and OCE 14595010 to MEF and DMG and, the National Oceanic and Atmospheric Administration award NA16SEC4810007 to DMG. We are grateful to the hardworking and professional crew of the R/V Savannah. Lee Ann DeLeo prepared the figures, Charles Y. Robertson proofread the manuscript and, James (Jimmy) Williams manufactured the plankton wheel

1. Preparing culturing facilities for rearing D. gegenbauri
NOTE: All materials and equipment required are listed in the Table of Materials.
<ol>
	<li>Prepare 1 M Sodium Hydroxide (NaOH), 0.06 M Potassium Permanganate (KMnO4) solution. To prepare this solution, dissolve 400 g of NaOH into 10 L deionized water. Add 100 g of KMnO4 to the NaOH solution and mix well.</li>
	<li>Prepare a 0.1 M Sodium bisulfite (NaHSO3) solution by dissolving 100 g of NaHSO3 into 10 L deionized water and mix well. 
	CAUTION: These reagents are irritants that may cause respiratory problems if inhaled. Place in a well-ventilated area such as a fume hood. Avoid any skin contact. Wear protective gloves, protective clothing, eye protection, and face protection when handling.</li>
	<li>Before establishing and rearing doliolid cultures in the laboratory, clean and sterilize the culture jars.
	<ol>
		<li>Rinse 1.9 L and 3.8 L culture jars at least 3 times with deionized water. Allow the screw caps to dry, as the caps are not included in the following cleaning steps.</li>
		<li>Clean and sterilize 1.9- and 3.8 L glass culture jars by immersing them in the NaOH/KMnO4 solution. Allow the jars to soak overnight.</li>
		<li>Remove the jars from the NaOH/KMnO4 solution and immerse the jars into the sodium bisulfite (NaHSO3) solution. Allow the jars to soak overnight.</li>
		<li>Remove the jars from the NaHSO3 solution and rinse thoroughly with deionized water. Allow the jars to dry.</li>
	</ol>
	</li>
	<li>Place the plankton wheel (Figure 2) in a temperature-controlled space (environmental chamber). Equilibrate the temperature to 20 &#176;C. For a more detailed description of the custom plankton wheel please refer to the Supplementary Figure 1.</li>
</ol>
2. Phytoplankton culture
<ol>
	<li>Obtain algal cultures from the National Center for Marine Algae and Microbiota (NCMA) or other sources to be used as food for D. gegenbauri. Mixtures of two flagellate species including Isochrysis galbana (CCMP 1323), Rhodomonas sp (CCMP 740), and a small diatom, Thalassiosira weissflogii (CCMP 1051) were obtained from the NCMA and have been used in previous laboratory studies to rear doliolids successfully17.</li>
	<li>Prepare L1 and L1-Si growth media22 as recommended by the NCMA.</li>
	<li>Follow the instructions provided by the supplier to initiate the new algal cultures.</li>
	<li>To maintain stock cultures, using rigorous axenic culture techniques, transfer 0.5 mL of old senescing culture to 25 mL of fresh growth media in&#160;sterile 55 mL glass culture tubes every two weeks. 
	NOTE: It is not possible to store living algal cultures without transferring them regularly. If cultures will not be used for long periods, and it is not possible to maintain cultures for the duration of the non-use period, it is recommended re-acquiring these common algal cultures from their original sources (e.g., NCMA).</li>
	<li>Prepare larger volumes of phytoplankton for feeding doliolids in clean 500 mL plastic tissue culture flasks containing 200 mL of growth media.
	<ol>
		<li>Inoculate phytoplankton from axenic stocks (4 mL) into 200 mL of growth media (1:50 dilution).</li>
		<li>Incubate at 20 &#176;C with a 12:12 h light:dark cycle under cool white light illumination of 65-85 &#181;E/m2. Lay culture flasks flat to maximize illumination. Gently swirl culture daily.</li>
		<li>Determine the concentration of cells using a particle counter or microscope to monitor the growth of the cultures. 
		NOTE: After 7-10 days from inoculation, the flagellate cultures will contain ~105-106 cells/mL and the diatom culture will contain ~104-105 cells/mL. These concentrations are enough to maintain the doliolid cultures.</li>
		<li>Initiate new feeding stocks at a minimum of every two weeks to provide enough algal biomass for supporting all culture activities.</li>
	</ol>
	</li>
</ol>
3. Collection of wild doliolids and seawater for culture
NOTE: An overview of collection and cultivation approaches is outlined in Figure 3. A description of the specialized collection plankton net and cod-end is provided in Figure 4.
<ol>
	<li>Locate doliolids by detecting them using either plankton nets or in situ imaging systems23. 
	NOTE: Because doliolids are rarely present in surface waters and are not detectable by remote sensing technology, guided by prerequisite knowledge of conditions favorable to doliolids (see Discussion), the presence of doliolids must be determined prior to sampling.</li>
	<li>Collect particle-rich seawater prior to collecting live doliolids in preparation for initiating a D. gegenbauri culture.
	<ol>
		<li>Deploy Niskin bottles mounted on a CTD rosette or equivalent equipment to collect water from the site where doliolids are located and from the depth containing the highest estimates of chlorophyll a concentration estimated by in situ fluorometry. 
		NOTE: Chlorophyll a concentration is used as an indicator of particle concentrations. On the South Atlantic Bight (SAB) mid-continental shelf, the subsurface chlorophyll a maximum is usually close to the bottom, but in other locations, it may not be.</li>
	</ol>
	</li>
	<li>Once doliolids are located, recover undamaged doliolid zooids using the specialized plankton net and cod-end. Before deploying the net, fill the cod-end with seawater.
	<ol>
		<li>From a drifting ship, lower and raise the net through the water column maintaining an oblique towing angle of ~15 - 25&#176; and vertical deployment and retrieval speed not greater than 15 m/min.</li>
	</ol>
	</li>
	<li>Once the net is onboard, gently transfer and divide the contents of the cod-end into 3, 5-gallon (~20 L) plastic buckets each containing ~ 10 L of surface seawater collected from the site. 
	NOTE: New plastic buckets should be conditioned by the addition of seawater days before living doliolid collection. The aim is to reduce the leaching of chemicals from plastic. If seawater is not available, use purified (e.g., Milli Q) or tap water free of toxic contaminants to condition the buckets.</li>
	<li>Isolate doliolid zooids from other plankton.
	<ol>
		<li>In small batches (~ 2 L) transfer mixed planktons from the net tow contents (now in 20 L plastic buckets) to a 2 L glass beaker.</li>
		<li>Using a wide-bore glass pipette (8 mm ID x 38 cm length), carefully siphon and transfer actively swimming doliolid zooids from the beaker into clean glass culture jars containing particle-rich seawater collected using Niskin bottles from where doliolids were located.</li>
		<li>Gently release the doliolid zooids beneath the surface of the seawater. 
		NOTE: Collect gonozooids, phorozooids containing attached developing gonozooids, and nurse stages containing attached trophozooids (Figure 1).</li>
	</ol>
	</li>
	<li>After the addition of doliolids, add Rhodomonas sp. culture to a final concentration of ~ 5 x 103 &#8211; 104 cells/mL (~50 mL of a culture containing ~ 5 x 104 &#8211; 1 x 105 cells/mL in a 3.8 L jar). This is to determine if the doliolids are actively feeding. When doliolids ingest Rhodomonas sp., their digestive tract will appear red in color. Remove zooids that do not appear to be feeding.</li>
	<li>To prevent doliolids from being trapped at the air-water interface, avoid the headspace in the culture jars by completely filling the jars with unfiltered particle-rich seawater and placing a piece of plastic wrap over the jar opening (89 mm wide).
	<ol>
		<li>Avoid creating air bubbles that can also damage the animals. Carefully screw the cap onto the jar and gently invert the jar to determine if bubbles are present. If bubbles are present, remove them.</li>
		<li>After jars are filled, wipe the excess water from the outside of the jar.</li>
	</ol>
	</li>
	<li>Mount each jar onto the plankton wheel (Figure 2) by placing the jar on the vertical metal bars covered with rubber tubing, and between a stainless-steel hose clamp.
	<ol>
		<li>Ensure that the back of the jar is cushioned against the rubber tubing. Tighten the hose clamp around the jar by adjusting the screw.</li>
		<li>Check that the jar is not moving once it is securely fastened in place. Allow the jars to rotate at 0.3 rpm to keep the doliolids in suspension. 
		CAUTION: It is important not to overtighten the jar to prevent the jar from cracking.</li>
	</ol>
	</li>
	<li>On the ship, maintain the culture vessels on the plankton wheel at 20 &#176;C in dim light until they can be transferred to the laboratory culture facility.</li>
	<li>Upon returning to the laboratory, transfer the jars containing doliolids into the prepared culture facility. Mount jars on the plankton wheel (see step 3.8) and allow the jars to continue to rotate at 0.3 rpm. 
	NOTE: All rearing of doliolids in this study was conducted at 20 &#176;C.</li>
</ol>
4. Maintaining D. gegenbauri cultures
<ol>
	<li>From the ship to the lab, allow the animals to acclimate in the original jars to the laboratory conditions for 3 days.
	<ol>
		<li>During the acclimation period, use a wide bore glass pipette to exchange 10% of the water with unfiltered particle-rich seawater from the collection site every day for 3 days.</li>
		<li>Keep several copepods in the jar but remove all other zooplankton, large fecal pellets, and large aggregated particles that may clog the doliolid&#8217;s filtering apparatus (mucus net). If the culture consists of early nurses, keep one large gonozooid&#160;(&#8805; 6 mm) in the jar. 
		NOTE: It is not important which copepod species are included in the culture, but in this experiment, the most abundant species present from where the doliolids were captured was used.</li>
	</ol>
	</li>
	<li>Following the acclimation period, transfer doliolid zooids and copepods from the original jar to a clean cultivation jar containing 80% glass fiber filter (GF/F) filtered seawater and 20% of the seawater from the original jar. Prepare filtered seawater by filtering seawater through a GF/F with a nominal pore size of 0.7 &#181;m filter paper.</li>
	<li>Maintain the new culture by exchanging 10% of the water with GF/F filtered seawater every 3 days and by removing aggregates and fecal pellets. Weekly, transfer animals to a new jar as described in step 4.2.</li>
	<li>Feed doliolids by maintaining phytoplankton concentrations in the culture jars between 40 - 95 &#181;g C/L. 
	NOTE: These concentrations mimic environmental conditions that are known to support bloom conditions for D. gegenbauri17. The mixture of algal species varies depending on the life stage and the number of zooids in each jar. During early life stages, add 1:1 mixture (by carbon content) of the cryptomonad algae (Isochrysis galbana and Rhodomonas sp.) only. Larger prey species can easily clog the feeding apparatus of small nurses and developing trophozooids. Add the diatom Thalassiosira weissflogii to the algal mixture, also at equal carbon content, when feeding larger nurses, phorozooids, and gonozooids.
	<ol>
		<li>Monitor algal concentrations pre- and post-feeding to guide the decision of how frequently and how much algae to add to the cultures. Use a particle counter to determine algal concentrations, because algal concentrations in the culture jars are relatively dilute.</li>
	</ol>
	</li>
	<li>Remove enough zooids to maintain algal concentrations of 40 &#8211; 95 &#181;gC/L so that the remaining doliolids will have enough food to grow. 
	NOTE: The most difficult life stage to maintain successfully under laboratory conditions is the developing larvae and oozooid (early nurse). During this phase of the culture, keep one large gonozooid (&#8805; 6 mm) in addition to several copepods in the jar with developing larvae and oozooids (~ 20 per 3.8 L jar).</li>
	<li>Transfer at least 4 nurses into to a new culturing jar once a minimum of 8 trophozooids are visible on the nurse&#8217;s cadophore (Figure 1B). 
	NOTE: Trophozooids will double in number every 1 &#8211; 2 days at 20 &#176;C. Trophozooids are large enough to be visible to the naked eye.
	<ol>
		<li>Remove two of the nurses once nurses develop 20 trophozooids.</li>
		<li>Remove one nurse when the nurses develop &#62; 30 trophozooids on their cadophores. Allow the remaining nurse to develop phorozooids on its cadophore.</li>
		<li>Remove the nurse once the nurse releases up to 30 phorozooids.</li>
	</ol>
	</li>
	<li>Reduce the number of animals in the jar once the phorozooids reach 3 mm in size.
	<ol>
		<li>Remove all but four phorozooids when the phorozooids become larger (&#62; 5 mm) and have developed gonozooid clusters.</li>
		<li>Reduce the culture to two phorozooids when the number of gonozooids clusters increase in size and begin to feed.</li>
		<li>Remove the phorozooids once the phorozooids release up to 30 gonozooids.</li>
	</ol>
	</li>
	<li>Reduce the number of gonozooids from 30 zooids to 2 per jar. Allow fertilized eggs to be released into the jar.
	<ol>
		<li>Remove one gonozooid leaving a single gonozooid in the jar once the oozooids develop. 
		NOTE: Discarded nurses, phorozooids, and gonozooids can be used to seed additional cultures and to conduct further experiments.</li>
	</ol>
	</li>
</ol>

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M., Paffenh&#246;fer, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Feeding+and+growth+rates+of+the+doliolid,+Dolioletta+gegenbauri+Uljanin+(Tunicata,+Thaliacea).">Feeding and growth rates of the doliolid, Dolioletta gegenbauri Uljanin (Tunicata, Thaliacea).</a> Journal of Plankton Research. 22 (8), 1485-1500 (2000).</li><li>Haskell, A., Hofmann, E., Paffenh&#246;fer, G., Verity, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modeling+the+effects+of+doliolids+on+the+plankton+community+structure+of+the+southeastern+US+continental+shelf.">Modeling the effects of doliolids on the plankton community structure of the southeastern US continental shelf.</a> Journal of Plankton Research. 21 (9), 1725-1752 (1999).</li><li>Gibson, D. M., Paffenh&#246;ffer, G. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Asexual+reproduction+of+the+doliolid,+Dolioletta+gegenbauri+Uljanin+(Tunicata,+Thaliacea).">Asexual reproduction of the doliolid, Dolioletta gegenbauri Uljanin (Tunicata, Thaliacea).</a> Journal of Plankton Research. 24 (7), 703-712 (2002).</li><li>Frischer, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reliability+of+qPCR+for+quantitative+gut+content+estimation+in+the+circumglobally+abundant+pelagic+tunicate+Dolioletta+gegenbauri+(Tunicata,+Thaliacea).">Reliability of qPCR for quantitative gut content estimation in the circumglobally abundant pelagic tunicate Dolioletta gegenbauri (Tunicata, Thaliacea).</a> Food Webs. 1, 7 (2014).</li><li>K&#246;ster, M., Paffenh&#246;fer, G., Baker, C., Williams, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oxygen+consumption+of+doliolids+(Tunicata,+Thaliacea).">Oxygen consumption of doliolids (Tunicata, Thaliacea).</a> Journal of Plankton Research. 32 (2), 171-180 (2010).</li><li>Paffenh&#246;fer, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+hypothesis+on+the+fate+of+blooms+of+doliolids+(Tunicata,+Thaliacea).">A hypothesis on the fate of blooms of doliolids (Tunicata, Thaliacea).</a> Journal of Plankton Research. 35 (4), 919-924 (2013).</li><li>Takahashi, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sapphirinid+copepods+as+predators+of+doliolids:+Their+role+in+doliolid+mortality+and+sinking+flux.">Sapphirinid copepods as predators of doliolids: Their role in doliolid mortality and sinking flux.</a> Limnology and Oceanography. 58 (6), 1972-1984 (2013).</li><li>Mart&#237;-Solans, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Oikopleura+dioica+Culturing+Made+Easy:+A+Low-Cost+Facility+for+an+Emerging+Animal+Model+in+EvoDevo.">Oikopleura dioica Culturing Made Easy: A Low-Cost Facility for an Emerging Animal Model in EvoDevo.</a> Genesis. 53 (1), 183-193 (2015).</li><li>Nishida, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+the+appendicularian+Oikopleura+dioica:+Culture,+genome,+and+cell+lineages.">Development of the appendicularian Oikopleura dioica: Culture, genome, and cell lineages.</a> Development Growth &amp; Differentiation. 50, S239-S256 (2008).</li></ol>

The authors have nothing to declare.

The capacity to culture doliolids has been established over the past several decades and has been used to support research in several areas. Experimental studies in our laboratories have supported the publication of at least 15 scientific studies focused on the feeding and growth18,26, reproduction18,28, diet6,29, physiology30</s...

Zooplankton account for the largest animal biomass in the ocean, are key components in marine food webs, and play important roles in ocean biogeochemical cycles1,2. Zooplankton, although comprised of a huge diversity of organisms, can be grossly distinguished into two categories: gelatinous and non-gelatinous with few intermediate taxa3,4. Compared to the non-gelatinous zooplankton, gelatinous zooplankton are especially difficult to study because of their complex life histories5, and their delicate tissues are easily damaged during capture and handling. Gelatinous zooplankton species are, therefore, notoriously difficult to culture in the laboratory and generally less studied compared to non-gelatinous species6.
Among gelatinous zooplankton groups, one abundant and of ecological importance in the world&#8217;s ocean are the Thaliaceans. Thaliaceans are a class of pelagic tunicates that include the orders Salpida, Pyrosomida, and Doliolida7. Doliolida, collectively referred to as doliolids, are small barrel-shaped free-swimming pelagic organisms that can reach high abundances in productive neritic regions of subtropical oceans. Doliolids are among the most abundant of all the zooplankton groups4,8. As suspension feeders, doliolids collect food particles from the water column by creating filter currents and capturing them on mucus nets9. Taxonomically, doliolids are classified in the phylum Urochordata10. Ancestral to the chordates, and in addition to their ecological significance as key components of marine pelagic systems, Thaliaceans are of significance to understanding the origins of colonial life history10,11 and the evolution of the chordates5,7,10,12,13,14.
The life history of doliolids is complex and contributes to the difficulty in culturing and sustaining them through their life cycle. A review of the doliolid life cycle and anatomy can be found in Godeaux et al.15. The doliolid life cycle, which involves an obligatory alternation between sexual and asexual life-history stages, is presented in Figure 1. Eggs and sperm are produced by the hermaphroditic gonozooids, the only solitary stage of the life cycle. Gonozooids release sperm to the water column and eggs are internally fertilized and released to develop into larvae. Larvae hatch and metamorphose into oozooids that can reach 1-2 mm. Presuming conducive environmental conditions and nutrition, oozooids become early nurses within 1-2 days at 20 &#176;C and initiate the colonial stages of the life cycle. Oozooids asexually produce buds on their ventral stolon. These buds leave the stolon and migrate to the dorsal cadophore where they line up in three paired rows. The central double rows become phorozooids and the outer two double rows become trophozooids. The latter provide food to both the nurse and the phorozooids16,17. The trophozooids supply the nurse with nutrition as she loses all internal organs. As the abundance of trophozooids increases, the size of the nurse can reach 15 mm in the laboratory. As the phorozooids grow, they increasingly ingest planktonic prey and reach ~ 1.5 mm in size prior to being released as individuals17. A single nurse may release &#62; 100 phorozooids during its lifespan18. After the phorozooids are released from the cadophore, they continue to grow and are the second colonial stage of the life cycle. Once they reach ~ 5 mm in size, each phorozooid develops a cluster of gonozooids on their ventral peduncle. These gonozooids can ingest particles when they reach ~1 mm in length. After the gonozooids have reached ~ 2 to 3 mm in size they are released from the phorozooid and become the only solitary stage of the life cycle. Once they reach ~ 6 mm in size, gonozooids become sexually mature17. Gonozooids can reach 9 mm or greater in length. Gonozooids are hermaphroditic, sperm is released intermittently while the fertilization of the eggs occurs internally16,17. When the gonozooid is &#8805; 6 mm in size, it releases up to 6 fertilized eggs. Successful culturing requires supporting the specific needs of each of these unique life history stages.
Due to the ecological and evolutionary significance of Thaliaceans, including doliolids, there is a need for the cultivation methodologies to advance the understanding of this organism&#8217;s unique biology, physiology, ecology, and evolutionary history19. Doliolids have considerable promise as experimental model organisms in developmental biology and functional genomics because they are transparent and likely have streamlined genomes20,21. The lack of reliable cultivation methods, however, impedes their usefulness as laboratory models. Although a handful of laboratories have published results based on cultured doliolids, to our knowledge cultivation approaches and detailed protocols have not been previously published. Based on years of experience, and trial and error cultivation attempts, the purpose of this study was to review experiences and to share protocols for the collection and cultivation of doliolids, specifically the species Dolioletta gegenbauri.

<table border="0" cellpadding="0" cellspacing="0" style="width:1860px;" width="1860">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Algal culture tubes (55 mL sterile disposable glass culture tubes)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td style="width:595px;">For algal cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Autoclave</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For sterilizing equipment and seawater for algal cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Beakers (2 L glass)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For sorting diluted plankton net tow contents</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Buckets (5 gallon, ~20L)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For diluting contents of planton net tow - should be seawater conditioned before first use</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Carboys (20 L)&#160;</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For storing seawater</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Doliolid glass culturing jar (1.9 L narrow mouth glass jar with cap)</td>
			<td>Qorpak</td>
			<td>GLC-01882</td>
			<td>Container for culture</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Doliolid glass culturing jar (3.8 L narrow mouth glass jar with cap)</td>
			<td>Qorpak</td>
			<td>GLC-01858</td>
			<td>Container for culture</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Environmental Chamber (Temperature controlled enviromental chamber)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>To accommodate plankton wheel and culture maintenance</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Filtration apparatus for 47 mm filters</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For filtering seawater for cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass microfiber filters, 47 mm</td>
			<td>Whatman</td>
			<td>1825-047</td>
			<td>For filtering seawater for cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Glass pipette (borosillicate glass pipette (glass tubing), OD 10mm, ID 8 mm, wall thickness 1mm)</td>
			<td>Science Company</td>
			<td>NC-10894</td>
			<td>Custom cut and edges polished</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Hose clamps, stainless steel, #104 (178 mm)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For holding culturing jars to the plankton wheel</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isochrysis galbana strain CCMP1323</td>
			<td>National Center for Marine Algae and Microbiota (NCMA)</td>
			<td>strain CCMP1323</td>
			<td>For feeding doliolid cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">L1 Media Kit, 50 L</td>
			<td>National Center for Marine Algae and Microbiota (NCMA)</td>
			<td>MKL150L</td>
			<td>For culturing algae</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Lamp (Fluorescent table lamp with an adjustable arm)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For illuminating doliolids in the jars and beakers</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Lighted temperature controlled incubator</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For algal cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Micropipettes and sterile tips (0-20 &#181;l, 20-200 &#181;l, 200-1000&#160;&#181;l)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For algal cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Plankton Net (202 &#181;m 0.5 m, 5:1 length) with cod end ring and &#160;4 L aquarium cod-end</td>
			<td>Sea-Gear Corporation</td>
			<td>90-50x5-200-4A/BB</td>
			<td>For collecting living doliolids (see Figure 4)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Plankton Wheel</td>
			<td style="width:417px;">NA</td>
			<td style="width:167px;">NA</td>
			<td>Custom built (see Figure 2)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Plastic wrap</td>
			<td>Any</td>
			<td>NA</td>
			<td>To cover inside of lid of doliolid culture jars</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Potassium Permanganate</td>
			<td>Fisher Scientific</td>
			<td>P279-500</td>
			<td>Reagent for cleaning jars and glassware</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rhodomonas&#160;sp. strain CCMP740</td>
			<td>National Center for Marine Algae and Microbiota (NCMA)</td>
			<td>strain CCMP740</td>
			<td>For feeding doliolid cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Rubber Tubing</td>
			<td style="width:417px;">NA</td>
			<td style="width:167px;">NA</td>
			<td>For holding culturing jars to the plankton wheel (can be made from tygon tubing)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Bisulfite</td>
			<td>Fisher Scientific</td>
			<td>S654-500</td>
			<td>Reagent for cleaning jars and glassware</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium Hydroxide</td>
			<td>Fisher Scientific</td>
			<td>BP359-212</td>
			<td>Reagent for cleaning jars and glassware</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Sterile serological pipettes (1 mL, 5 mL, 10 mL, 25 mL)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For algal cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Thalassiosira weissflogii strain CCMP1051</td>
			<td>National Center for Marine Algae and Microbiota (NCMA)</td>
			<td>strain CCMP1051</td>
			<td>For feeding doliolid cultures</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:681px;">Tissue culture flasks (250 mL)</td>
			<td style="width:417px;">Any</td>
			<td style="width:167px;">NA</td>
			<td>For algal cultures</td>
		</tr>
	</tbody>
</table>

cultivation of the marine pelagic tunicate dolioletta gegenbauri (uljanin 1884) for experimental studies

Gelatinous zooplanktons play a crucial role in ocean ecosystems. However, it is generally difficult to investigate their physiology, growth, fecundity, and trophic interactions primarily due to methodological challenges, including the ability to culture them. This is particularly true for the doliolid, Dolioletta gegenbauri. D. gegenbauri commonly occurs in productive subtropical continental shelf systems worldwide, often at bloom concentrations capable of consuming a large fraction of daily primary production. In this study, we describe cultivation approaches for collecting, rearing, and maintaining D. gegenbauri for the purpose of conducting laboratory-based studies. D. gegenbauri and other doliolid species can be captured live using obliquely towed conical 202 &micro;m mesh plankton nets from a drifting ship. Cultures are most reliably established when water temperatures are below 21 &deg;C and are started from immature gonozooids, maturing phorozooids, and large nurses. Cultures can be maintained in rounded culture vessels on a slowly rotating plankton wheel and sustained on a diet of cultured algae in natural seawater for many generations. In addition to the ability to establish laboratory cultures of D. gegenbauri, we demonstrate that the collection condition, algae concentration, temperature, and exposure to naturally conditioned seawater are all critical to the culture establishment, growth, survival, and reproduction of D. gegenbauri.

Following the described procedures for collecting and culturing the doliolid, D. gegenbauri outlined in Figure 3, it is possible to maintain a culture of D. gegenbauri throughout its complex life history (Figure 1) and sustain it for many generations. Although cultivation of D. gegenbauri is described here, these procedures should also be relevant for the cultivation of other doliolid species.
Capturing heal...

Watch this Scientific Journal Video about Cultivation of the Marine Pelagic Tunicate Dolioletta gegenbauri Uljanin 1884 for Experimental Studies at JoVE.com

Cultivation of the Marine Pelagic Tunicate Dolioletta gegenbauri Uljanin 1884 for Experimental Studies

Doliolids, including the species Dolioletta gegenbauri, are small gelatinous marine zooplankton of ecological significance found on productive subcontinental shelf systems worldwide. The difficulty of culturing these delicate organisms limits their investigation. In this study, we describe cultivation approaches for collecting, rearing, and maintaining the doliolid Dolioletta gegenbauri.

Cultivation of the Marine Pelagic Tunicate Dolioletta gegenbauri (Uljanin 1884) for Experimental Studies

Gelatinous zooplanktons play a crucial role in ocean ecosystems. However, it is generally difficult to investigate their physiology, growth, fecundity, ...

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9.4K Views.  University of Georgia. Doliolids, including the species Dolioletta gegenbauri, are small gelatinous marine zooplankton of ecological significance found on productive subcontinental shelf systems worldwide. The difficulty of culturing these delicate organisms limits their investigation. In this study, we describe cultivation approaches for collecting, rearing, and maintaining the doliolid Dolioletta gegenbauri.

Video: Cultivation of the Marine Pelagic Tunicate Dolioletta gegenbauri Uljanin 1884 for Experimental Studies

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 ...

Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination control. Developments of some techniques aiming to reduce trace metal contamination in the last couple of decades have resulted in concentrations reported now being orders of magnitude lower than those in the past. These low concentrations often necessitate preconcentration of water samples prior to instrumental analysis of samples. Since contamination can appear in all phases of trace metal analyses, including sample collection (and during preparation of sampling containers), storage and handling, pretreatments, and instrumental analysis, specific care needs to be taken in order to reduce contamination levels at all steps. The effort to develop and utilize "clean techniques" in trace metal studies allows scientists to investigate trace metal distributions and chemical and biological behavior in greater details. This advancement also provides the required accuracy and precision of trace metal data allowing for environmental conditions to be related to trace metal concentrations in aquatic environments.
This protocol that is presented here details needed materials for sample preparation, sample collection, sample pretreatment including preconcentration, and instrumental analysis. By reducing contamination throughout all phases mentioned above for trace metal analysis, much lower detection limits and thus accuracy can be achieved. The effectiveness of "clean techniques" is further demonstrated using low field blanks and good recoveries for standard reference material. The data quality that can be obtained thus enables the assessment of trace metal distributions and their relationships to environmental parameters.

Special care using "clean techniques" is required to properly collect and process water samples for trace metal studies in aquatic environments. A protocol for sampling, processing, and analytical procedures with the aim of obtaining reliable environmental monitoring data and results with high sensitivity for detailed trace metal studies is presented.

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination ...

Experimental Protocol for Biodiesel Production with Isolation of Alkenones as Coproducts from Commercial Isochrysis Algal Biomass

The need to replace petroleum fuels with alternatives from renewable and more environmentally sustainable sources is of growing importance. Biomass-derived biofuels have gained considerable attention in this regard, however first generation biofuels from edible crops like corn ethanol or soybean biodiesel have generally fallen out of favor. There is thus great interest in the development of methods for the production of liquid fuels from domestic and superior non-edible sources. Here we describe a detailed procedure for the production of a purified biodiesel from the marine microalgae Isochrysis. Additionally, a unique suite of lipids known as polyunsaturated long-chain alkenones are isolated in parallel as potentially valuable coproducts to offset the cost of biodiesel production. Multi-kilogram quantities of Isochrysis are purchased from two commercial sources, one as a wet paste (80% water) that is first dried prior to processing, and the other a dry milled powder (95% dry). Lipids are extracted with hexanes in a Soxhlet apparatus to produce an algal oil (&#34;hexane algal oil&#34;) containing both traditional fats (i.e., triglycerides, 46-60% w/w) and alkenones (16-25% w/w). Saponification of the triglycerides in the algal oil allows for separation of the resulting free fatty acids (FFAs) from alkenone-containing neutral lipids. FFAs are then converted to biodiesel (i.e., fatty acid methyl esters, FAMEs) by acid-catalyzed esterification while alkenones are isolated and purified from the neutral lipids by crystallization. We demonstrate that biodiesel from both commercial Isochrysis biomasses have similar but not identical FAME profiles, characterized by elevated polyunsaturated fatty acid contents (approximately 40% w/w). Yields of biodiesel were consistently higher when starting from the Isochrysis wet paste (12% w/w vs. 7% w/w), which can be traced to lower amounts of hexane algal oil obtained from the powdered Isochrysis product.

Experimental Protocol for Biodiesel Production with Isolation of Alkenones as Coproducts from Commercial Isochrysis Algal Biomass

Detailed methods are presented for the production of biodiesel along with the co-isolation of alkenones as valuable coproducts from commercial Isochrysis microalgae.

The need to replace petroleum fuels with alternatives from renewable and more environmentally sustainable sources is of growing importance. ...

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 ...

Vegetated Treatment Systems for Removing Contaminants Associated with Surface Water Toxicity in Agriculture and Urban Runoff

Urban stormwater and agriculture irrigation runoff contain a complex mixture of contaminants that are often toxic to adjacent receiving waters. Runoff may be treated with simple systems designed to promote sorption of contaminants to vegetation and soils and promote infiltration. Two example systems are described: a bioswale treatment system for urban stormwater treatment, and a vegetated drainage ditch for treating agriculture irrigation runoff. Both have similar attributes that reduce contaminant loading in runoff: vegetation that results in sorption of the contaminants to the soil and plant surfaces, and water infiltration. These systems may also include the integration of granulated activated carbon as a polishing step to remove residual contaminants. Implementation of these systems in agriculture and urban watersheds requires system monitoring to verify treatment efficacy. This includes chemical monitoring for specific contaminants responsible for toxicity. The current paper emphasizes monitoring of current use pesticides since these are responsible for surface water toxicity to aquatic invertebrates.

This article summarizes the design attributes and the effectiveness of treatment systems that treat urban stormwater and agriculture irrigation runoff to remove pesticides and other contaminants associated with aquatic toxicity.

Urban stormwater and agriculture irrigation runoff contain a complex mixture of contaminants that are often toxic to adjacent receiving waters. Runoff may ...

Extraction of Organochlorine Pesticides from Plastic Pellets and Plastic Type Analysis

Plastic resin pellets, categorized as microplastics (&#8804;5 mm in diameter), are small granules that can be unintentionally released to the environment during manufacturing and transport. Because of their environmental persistence, they are widely distributed in the oceans and on beaches all over the world. They can act as a vector of potentially toxic organic compounds (e.g., polychlorinated biphenyls) and might consequently negatively affect marine organisms. Their possible impacts along the food chain are not yet well understood. In order to assess the hazards associated with the occurrence of plastic pellets in the marine environment, it is necessary to develop methodologies that allow for rapid determination of associated organic contaminant levels. The present protocol describes the different steps required for sampling resin pellets, analyzing adsorbed organochlorine pesticides (OCPs) and identifying the plastic type. The focus is on the extraction of OCPs from plastic pellets by means of a pressurized fluid extractor (PFE) and on the polymer chemical analysis applying Fourier Transform-InfraRed (FT-IR) spectroscopy. The developed methodology focuses on 11 OCPs and related compounds, including dichlorodiphenyltrichloroethane (DDT) and its two main metabolites, lindane and two production isomers, as well as the two biologically active isomers of technical endosulfan. This protocol constitutes a simple and rapid alternative to existing methodology for evaluating the concentration of organic contaminants adsorbed on plastic pieces.

Microplastics act as vector of potentially toxic organic contaminants with unpredictable effects. This protocol describes an alternative methodology for assessing the levels of organochlorine pesticides adsorbed on plastic pellets and identifying the polymer chemical structure. The focus is on pressurized fluid extraction and attenuated total reflectance Fourier transform infrared spectroscopy.

Plastic resin pellets, categorized as microplastics (&#8804;5 mm in diameter), are small granules that can be unintentionally released to the environment ...

Development of New Methods for Quantifying Fish Density Using Underwater Stereo-video Tools

The use of video camera systems in ecological studies of fish continues to gain traction as a viable, non-extractive method of measuring fish lengths and estimating fish abundance. We developed and implemented a rotating stereo-video camera tool that covers a full 360 degrees of sampling, which maximizes sampling effort compared to stationary camera tools. A variety of studies have detailed the ability of static, stereo-camera systems to obtain highly accurate and precise measurements of fish; the focus here was on the development of methodological approaches to quantify fish density using rotating camera systems. The first approach was to develop a modification of the metric MaxN, which typically is a conservative count of the minimum number of fish observed on a given camera survey. We redefine MaxN to be the maximum number of fish observed in any given rotation of the camera system. When precautions are taken to avoid double counting, this method for MaxN may more accurately reflect true abundance than that obtained from a fixed camera. Secondly, because stereo-video allows fish to be mapped in three-dimensional space, precise estimates of the distance-from-camera can be obtained for each fish. By using the 95% percentile of the observed distance from camera to establish species-specific areas surveyed, we account for differences in detectability among species while avoiding diluting density estimates by using the maximum distance a species was observed. Accounting for this range of detectability is critical to accurately estimate fish abundances. This methodology will facilitate the integration of rotating stereo-video tools in both applied science and management contexts.

We describe a new method for counting fishes, and estimating relative abundance (MaxN) and fish density using rotating stereo-video camera systems. We also demonstrate how to use distance from camera (Z distance) to estimate species-specific detectability.

The use of video camera systems in ecological studies of fish continues to gain traction as a viable, non-extractive method of measuring fish lengths and ...

Methods for Image-based Surveys of Benthic Macroinvertebrates and Their Habitat Exemplified by the Drop Camera Survey for the Atlantic Sea Scallop

Underwater imaging has long been used in the field of marine ecology but decreasing costs of high-resolution cameras and data storage have made the approach more practical than in the past. Image-based surveys allow for initial samples to be revisited and are non-invasive compared to traditional survey methods that typically involve nets or dredges. Protocols for image-based surveys can vary greatly but should be driven by target species behavior and survey objectives. To demonstrate this, we describe our most recent methods for an Atlantic sea scallop (Placopecten magellanicus) drop camera survey to provide a procedural example and representative results. The procedure is divided into three critical steps that include survey design, data collection, and data products. The influence of scallop behavior and the survey goal of providing an independent assessment of the U.S. sea scallop resource on the survey procedure are then discussed in the context of generalizing the method. Overall, the broad applicability and flexibility of the University of Massachusetts Dartmouth School for Marine Science and Technology (SMAST) drop camera survey demonstrates the method could be generalized and applied to a variety of sessile invertebrates or habitat focused research.

Image based surveying is an increasingly practical, non-invasive method to sample the marine environment. We present the protocol of a drop camera survey that estimates the abundance and distribution of the Atlantic sea scallop (Placopecten magellanicus). We discuss how this protocol can be generalized for application to other benthic macroinvertebrates.

Underwater imaging has long been used in the field of marine ecology but decreasing costs of high-resolution cameras and data storage have made the ...