Initial development of the low-volume ventilated culture system was supported in part by the National Science Foundation (CRI-OA-1416690 to Dickinson College). Dr. Lauren Mullineaux kindly provided laboratory facilities at the Woods Hole Oceanographic Institution, where the data presented for this system (Figure 4) was collected.

1. Routine maneuvers for establishing and maintaining larval cultures of C. fornicata
NOTE: This method starts with a gallon (3.8 L) jar of seawater containing adult C. fornicata that have just released brooded veliger larvae. Adults may be field-collected or obtained from a supplier given in the Table of Materials. The adults are protandrous hermaphrodites that live in mating stacks with sessile brooding females at the bottom of the stack. Do not break up adult stacks. Seasonality of reproduction and methods for conditioning adults for spawning out of season have been previously described17. Larvae are best collected within 2-3 h after release when they are strongly geonegative and will concentrate near the surface of the jar.
<ol>
	<li>Make a sieve by cutting the bottom off of a 400 mL tri-corner disposable plastic beaker and gluing on a panel of 236 &#181;m nylon mesh. For this purpose and especially for mesocosm construction (below), use a hot melt glue that is formulated for good adhesion to polyethylene.</li>
	<li>Remove aeration from the adult jar and let it stand for 15-20 min so that debris settles and larvae can swim easily to the surface.</li>
	<li>Siphon the larvae through a short length (15-20 cm) of 5 mm glass tubing affixed to the end of a 60-80&#160;cm length of soft plastic aquarium tubing into the sieve prepared as above, suspended in a 600 mL glass beaker such that excess seawater overflows the glass beaker while larvae are retained on the sieve. Keep the bottom of the sieve underwater and avoid stranding larvae.</li>
	<li>Briefly lift the sieve out of the beaker and use a squirt bottle of filtered seawater (FSW) to rinse the larvae into a glass bowl of FSW that is of a convenient size to manipulate under a low-power dissecting microscope. 
	NOTE: Larvae of C. fornicata thrive in filtered natural seawater near 30 ppt salinity or Instant Ocean artificial seawater made up to 32 ppt. salinity, at 20-25 &#176;C17,18. Filtration at &#60; 1 &#181;m is sufficient to eliminate fouling epizootic protists that can cause problems in laboratory cultures.</li>
	<li>Manually transfer and count the desired number of larvae into a culture jar using a Pasteur pipet. For best results, use 1 larva/5 mL culture volume for C. fornicata. Count the larvae and avoid the temptation to estimate numbers.</li>
	<li>Feed larvae with the desired ration of microalgae when culture is started and every other day replace the microalgae when culture water is replaced. 
	NOTE: A diet of 1 x 105 cells/mL Isochrysis galbana (strain T-ISO) is a good standard diet that yields near-maximal laboratory growth rates. Food ration and methods for the culture of T-ISO are provided in an earlier publication17.</li>
	<li>Change culture water every other day by pouring the contents of the culture into a sieve within a beaker, as in step 1.3&#160;above. Lift out the sieve and use a squirt bottle of FSW to rinse larvae into a culture jar of fresh FSW with new food. 
	&#8203;NOTE: Best practice is to segregate any glassware used for larval culture and to avoid any contact of that glassware with fixatives, soluble heavy metal salts, or detergents. Routine cleaning is best done with a paste of baking soda (NaHCO3) and a nylon scrubbing pad, while mineral deposits can be removed with a mild acid wash in white vinegar or dilute HCl.</li>
</ol>
2. Construction of ventilated cultures for larvae of C. fornicata
NOTE: The recommended glass jar (Table of Materials) has a polypropylene lid, which is inert to seawater and has the right thickness for fastening tubing barb inlets for a ventilating gas stream.
<ol>
	<li>Drill two 5 mm holes in the lid of each culture jar (including the lid&#39;s inner liner), each about 1.5 cm from the edge and opposite each other. (A 13/64 inch imperial or #7 ASME standard machinist&#39;s drill bit will also make an appropriate clearance hole).</li>
	<li>Install a 1/8 inch x 10-32 threaded nylon tubing adapter in each hole, using a 10-32 nylon nut, with the tubing barb on the outer surface of the lid.</li>
	<li>Apply a dab of silicone rubber aquarium sealant on the aperture of the threaded inner part of one of the tubing adapters and push a 20 cm length of 2 mm (outer diameter) polyethylene tubing through the aperture from the inside so that it protrudes a few mm beyond the outer aperture of the tubing barb.</li>
	<li>The tubing will now have a small plug of aquarium sealant at the end that protrudes through the tubing barb, but clear tubing will be visible just beyond the end of the barb. Let the sealant cure, then trim the protruding end of the polyethylene tubing so that it is flush with the end of the tubing barb.</li>
	<li>Fill the culture jar with FSW to within 1 cm of the shoulder at the top of the jar, screw down the lid, and attach the ventilating air or experimental gas mix supply to the tubing barb that holds the polyethylene ventilation tubing. Trim the length of the ventilation tubing so that its end lies neatly at the bottom of the jar.</li>
	<li>Adjust the flow of the ventilating air or gas mix to yield a slow stream of bubbles (Figure 1). Use an aquarium air pump with common aquarium gang valves to supply ventilation with ambient air or use mass-flow controllers for other experimental atmospheric gas mixes14.</li>
</ol>
3. Construction of a field-deployable mesocosm culture for larvae of C. fornicata
<ol>
	<li>Cut 4 evenly spaced rectangular openings, each 25 cm x 14.5 cm, in the sides of a standard 7-gallon polyethylene bucket. Position the bottom of each opening about 3.5 cm from the inside bottom of the bucket (Figure 2A). A hand-held electric saber saw with a fine-toothed blade is ideal for this purpose. Keep the cut-out waste pieces and slice them lengthwise into 2 cm-wide strips with a table saw.</li>
	<li>Cut out 4 panels of 236 &#181;m nylon mesh, each at least 30 cm x 20 cm, so as to comfortably overlap (by at least 2 cm) the cutout openings in the bucket. Temporarily secure one edge of a mesh panel over a long edge of its cutout using small spring clamps or clothespins.</li>
	<li>Glue the edge of the mesh panel to the bucket opening with hot melt glue that is formulated for polyethylene. Once the first edge is fastened, glue down the other edges while maintaining tension to get a taut surface on the mesh panel.</li>
	<li>Trim the lengths of the strips of bucket waste pieces from step 3.1, to make reinforcing strips to neatly cover the glued area where the mesh panels overlap the bucket openings. Press each covering strip in place with a liberal amount of hot melt glue, working quickly to get each strip in place before the glue hardens.</li>
	<li>Secure the end of each strip with a nylon blind rivet installed from the outside of the bucket. Trim excess glue and mesh from the outside edges of the reinforcing strips with a utility razor knife. 
	NOTE: Each rivet requires a 15/64 inch pilot hole. The interior (blind) parts of the rivets are visible on the inside of the finished mesocosm in Figure 2B.</li>
	<li>Deploy the mesocosm in a floating rack that holds the top of the bucket well above water (Figure 2C). The floating rack that is shown is made from 30 cm segments of 2 inch Schedule 40 PVC pipe, cemented together with elbow and tee joints to make an airtight structure that can hold a row of 4 replicate mesocosms.</li>
	<li>Depending on local conditions, the mesh panels may become fouled in 1-2 days, in a way that impedes exchange of seawater through the mesocosm. To the extent that this occurs, transfer larvae to a fresh clean mesocosm by pulling the fouled mesocosm 2/3rd out of the water and repeatedly bailing from its bottom into the fresh mesocosm with a 2 L pitcher.</li>
	<li>Perform 20-25 iterations to transfer most of the larvae, or until larvae are no longer observed in the fouled mesocosm. Clean the fouled mesocosm by gently scrubbing the mesh panels with a soft sponge loaded with a paste of baking soda (NaHCO3) and rinse with tap water.</li>
</ol>

Culturing Larvae of Crepidula fornicata in Laboratory and Field Conditions

<ol>
	<li>Conklin, E. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+embryology+of+Crepidula:+a+contribution+to+the+cell+lineage+and+early+development+of+some+marine+gastropods.">The embryology of Crepidula: a contribution to the cell lineage and early development of some marine gastropods.</a> Gasteropods. , (1897).</li><li>Henry, J. J., Collin, R., Perry, K. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+slipper+snail,+Crepidula:+an+emerging+lophotrochozoan+model+system.">The slipper snail, Crepidula: an emerging lophotrochozoan model system.</a> Biol Bull. 218 (3), 211-229 (2010).</li><li>Lyons, D. C., Henry, J. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Slipper+snail+tales:+how+Crepidula+fornicata+and+Crepidula+atrasolea+became+model+molluscs.">Slipper snail tales: how Crepidula fornicata and Crepidula atrasolea became model molluscs.</a> Curr Top Dev Biol. 147, 375-399 (2022).</li><li>Blanchard, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spread+of+the+slipper+limpet+Crepidula+fornicata+(L.+1758)+in+Europe.+Current+state+and+consequences.">Spread of the slipper limpet Crepidula fornicata (L. 1758) in Europe. Current state and consequences.</a> ScientiaMarina. 61 (Suppl 2), 109-118 (1997).</li><li>Beninger, P., Valdizan, A., Decottigies, P., Cognie, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Field+reproductive+dynamics+of+the+invasive+slipper+limpet,+Crepidula+fornicata.">Field reproductive dynamics of the invasive slipper limpet, Crepidula fornicata.</a> J Exp Mar Biol Ecol. 390 (2), 179-187 (2010).</li><li>Pechenik, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+relationship+between+temperature,+growth+rate,+and+duration+of+planktonic+life+for+larvae+of+the+gastropod+Crepidula+fornicata+(L.).">The relationship between temperature, growth rate, and duration of planktonic life for larvae of the gastropod Crepidula fornicata (L.).</a> J Exp Mar Biol Ecol. 74 (3), 241-257 (1984).</li><li>Pechenik, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Latent+effects:+surprising+consequences+of+embryonic+and+larval+experience+on+life+after+metamorphosis.">Latent effects: surprising consequences of embryonic and larval experience on life after metamorphosis.</a> Evol Ecol Marine Invertebrate Larvae. , 208-225 (2018).</li><li>Pechenik, J. A., Gee, C. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Onset+of+metamorphic+competence+in+larvae+of+the+gastropod+Crepidula+fornicata+(L.),+judged+by+a+natural+and+an+artificial+cue.">Onset of metamorphic competence in larvae of the gastropod Crepidula fornicata (L.), judged by a natural and an artificial cue.</a> J Exp Mar Biol Ecol. 167 (1), 59-72 (1993).</li><li>Taris, M., Comtet, T., Viard, F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Inhibitory+function+of+nitric+oxide+on+the+onset+of+metamorphosis+in+competent+larvae+of+Crepidula+fornicata:+A+transcriptional+perspective.">Inhibitory function of nitric oxide on the onset of metamorphosis in competent larvae of Crepidula fornicata: A transcriptional perspective.</a> Mar Genomics. 2 (3-4), 161-167 (2009).</li><li>Penniman, J. R., Doll, M. K., Pires, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neural+correlates+of+settlement+in+veliger+larvae+of+the+gastropod,+Crepidula+fornicata.">Neural correlates of settlement in veliger larvae of the gastropod, Crepidula fornicata.</a> Invertebrate Biol. 132 (1), 14-26 (2013).</li><li>Meyer-Kaiser, K. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Carryover+effects+of+brooding+conditions+on+larvae+in+the+slipper+limpet+Crepidula+fornicata.">Carryover effects of brooding conditions on larvae in the slipper limpet Crepidula fornicata.</a> Mar Ecol Prog Ser. 643, 87-97 (2020).</li><li>Noisette, F., 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=Does+encapsulation+protect+embryos+from+the+effects+of+ocean+acidification?+The+example+of+Crepidula+fornicata.">Does encapsulation protect embryos from the effects of ocean acidification? The example of Crepidula fornicata.</a> PLoS One. 9 (3), e93021 (2014).</li><li>Kriefall, N. G., Pechenik, J. A., Pires, A., Davies, S. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Resilience+of+Atlantic+slippersnail+Crepidula+fornicata+larvae+in+the+face+of+severe+coastal+acidification.">Resilience of Atlantic slippersnail Crepidula fornicata larvae in the face of severe coastal acidification.</a> Front in Mar Sci. 5, 312 (2018).</li><li>Bogan, S. N., McMahon, J. B., Pechenik, J. A., Pires, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Legacy+of+multiple+stressors:+Responses+of+gastropod+larvae+and+juveniles+to+ocean+acidification+and+nutrition.">Legacy of multiple stressors: Responses of gastropod larvae and juveniles to ocean acidification and nutrition.</a> Biol Bull. 236 (3), 159-173 (2019).</li><li>Pechenik, J. A., 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=Impact+of+ocean+acidification+on+growth,+onset+of+competence,+and+perception+of+cues+for+metamorphosis+in+larvae+of+the+slippershell+snail,+Crepidula+fornicata.">Impact of ocean acidification on growth, onset of competence, and perception of cues for metamorphosis in larvae of the slippershell snail, Crepidula fornicata.</a> Mar Biol. 166, 128 (2019).</li><li>Reyes, C. L., 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=The+marine+gastropod+Crepidula+fornicata+remains+resilient+to+ocean+acidification+across+two+life+history+stages.">The marine gastropod Crepidula fornicata remains resilient to ocean acidification across two life history stages.</a> Front Physiol. 12, 702864 (2021).</li><li>Pires, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Artificial+seawater+culture+of+the+gastropod+Crepidula+fornicata+for+studies+of+larval+settlement+and+metamorphosis.+In:+Carroll,+D.,+Stricker,+S.+(eds)+Developmental+biology+of+the+sea+urchin+and+other+marine+invertebrates.">Artificial seawater culture of the gastropod Crepidula fornicata for studies of larval settlement and metamorphosis. In: Carroll, D., Stricker, S. (eds) Developmental biology of the sea urchin and other marine invertebrates.</a> Methods Mol Biol. 1128, 35-44 (2014).</li><li>Bashevkin, S. M., Pechenik, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+interactive+influence+of+temperature+and+salinity+on+larval+and+juvenile+growth+in+the+gastropod+Crepidula+fornicata+(L.).">The interactive influence of temperature and salinity on larval and juvenile growth in the gastropod Crepidula fornicata (L.).</a> J Exp Mar Biol Ecol. 470, 78-91 (2015).</li><li>Strathmann, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Reproduction and Development of Marine Invertebrates of the Northern Pacific Coast: Data and Methods for the Study of Eggs, Embryos, and Larvae. , (1987).</li><li>Clark, H. R., Gobler, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diurnal+fluctuations+in+CO2+and+dissolved+oxygen+concentrations+do+not+provide+a+refuge+from+hypoxia+and+acidification+for+early-life-stage+bivalves.">Diurnal fluctuations in CO2 and dissolved oxygen concentrations do not provide a refuge from hypoxia and acidification for early-life-stage bivalves.</a> Mar Ecol Prog Ser. 558, 43114 (2016).</li><li>Gobler, C. J., Baumann, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hypoxia+and+acidification+in+ocean+ecosystems:+coupled+dynamics+and+effects+on+marine+life.">Hypoxia and acidification in ocean ecosystems: coupled dynamics and effects on marine life.</a> Biol Lett. 12 (5), 20150976 (2016).</li><li>Pechenik, J. A., Hilbish, T. J., Eyster, L. S., Marshall, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relationship+between+larval+and+juvenile+growth+rates+in+two+marine+gastropods,+Crepidula+plana+and+C.">Relationship between larval and juvenile growth rates in two marine gastropods, Crepidula plana and C.</a> fornicata. Mar Biol. 125, 119-127 (1996).</li><li>Padilla, D. K., McCann, M. J., McCarty Glenn, M., Hooks, A. P., Shumway, S. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+food+on+metamorphic+competence+in+the+model+system+Crepidula+fornicata.">Effect of food on metamorphic competence in the model system Crepidula fornicata.</a> Biol Bull. 227 (3), 242-251 (2014).</li><li>Wallace, R. B., Baumann, H., Grear, J. S., Aller, R. C., Gobler, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Coastal+ocean+acidification:+the+other+eutrophication+problem.">Coastal ocean acidification: the other eutrophication problem.</a> Estuarine, Coast Shelf Sci. 148, 1-13 (2014).</li></ol>

There are no conflicts of interest to report.

Although larvae of C. fornicata are relatively easy to culture compared to other planktotrophic marine larvae, attention to fundamentals of good culture practice is still essential17,19. Healthy larvae should begin to feed immediately after hatching. This is easily verified on the day after hatching by observing their full guts, packed with algal cells, using a dissecting microscope with transillumination. Shells of healthy larvae should remain clean, br...

The slipper limpet, Crepidula fornicata (Gastropoda: Calyptraeidae), is well-represented in current and historical research literature because of its utility as a developmental model and because of its widespread impacts as an invasive species. It served as a foundational example of spiralian development in the classic age of experimental embryology1 and has experienced a rebirth of interest with the application of modern imaging and genomic tools to dissect mechanisms of lophotrochozoan early development2,3. At the other end of its life history, other investigations have focused on the impacts of adult populations of this ecosystem engineer in temperate coastal marine environments far removed from its original distribution in eastern North America4,5. In between embryo and adult, the veliger larvae of this species have been subjects of numerous studies of larval development and ecology, especially of factors influencing growth and acquisition of competence for metamorphosis, the internal and external cues mediating larval settlement, and the effects of larval experience on juvenile performance6,7,8,9,10,11. Recent studies have revealed the resilience of larvae and juveniles of C. fornicata to ocean acidification, yet another avenue for productive research use of this animal12,13,14,15,16.
An advantage of C. fornicata for studies of marine larval biology is that it is relatively easy to grow in the laboratory in natural or artificial seawater on a unialgal diet of the flagellate Isochrysis galbana. Culture methods have been detailed by the author in an earlier methods-focused print publication17. The reasons for the present contribution are twofold. First, the routine physical maneuvers involved in establishing and caring for cultures are conceptually very simple but difficult to perform correctly without hands-on or video demonstration. Second, two variations on previously described culture methods are described that are especially suited to laboratory and field studies of responses to environmental stressors such as ocean acidification, eutrophication, and oxygen depletion. The first of these is a low-volume (800 mL) culture system suited for manipulation of pH and dissolved oxygen in seawater via small volumes of bubbled gases, and the second is a larger volume (15 L) mesocosm system that can be placed in the field and that allows free exchange of ambient seawater.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Bucket, Polyethylene, 7 gallon</td>
			<td>US Plastic</td>
			<td>16916</td>
			<td>for mesocosm</td>
		</tr>
		<tr>
			<td>Crepidula fornicata</td>
			<td>Marine Biological Laboratory, Marine Resources Center</td>
			<td>760</td>
			<td>adult broodstock</td>
		</tr>
		<tr>
			<td>Hotmelt glue, Infinity Supertac 500</td>
			<td>Hotmelt.com</td>
			<td>INFINITY IM-SUPERTAC-500-12-1LB</td>
			<td>good for bonding polyethylene</td>
		</tr>
		<tr>
			<td>Jar, glass, 32 oz, with polypropylene lid</td>
			<td>Uline</td>
			<td>S-19316P-W</td>
			<td>for 800 mL ventilated cultures</td>
		</tr>
		<tr>
			<td>Nitex mesh, 236 &#181;m</td>
			<td>Dynamic Aqua Supply Ltd.</td>
			<td>NTX236-136</td>
			<td>for mesocosm</td>
		</tr>
		<tr>
			<td>Nut, hex, nylon, 10-32 thread</td>
			<td>Home Depot</td>
			<td>1004554441</td>
			<td>for fastening tubing barbs</td>
		</tr>
		<tr>
			<td>Rivets, nylon, blind, 15/64&#34; diameter, 5/32&#34;-5/16&#34; grip range, pack of 8</td>
			<td>NAPA auto parts</td>
			<td>BK 6652844</td>
			<td>4 packs needed per mesocosm</td>
		</tr>
		<tr>
			<td>Tubing barb 1/8&#34; x 10-32 thread</td>
			<td>US Plastic</td>
			<td>65593</td>
			<td>2 needed per culture jar</td>
		</tr>
		<tr>
			<td>Tubing, polyethylene, 2.08 mm OD</td>
			<td>Fisher Scientific</td>
			<td>14-170-11G</td>
			<td>for ventilating gas stream inside culture jar</td>
		</tr>
		<tr>
			<td>Tubing, Tygon, 1/8&#34;x3/16&#34;x1/32&#34;</td>
			<td>US Plastic</td>
			<td>57810</td>
			<td>fits barbs for ventilating cultures</td>
		</tr>
	</tbody>
</table>

laboratory and field culture of larvae of the slipper limpet, crepidula fornicata

The calyptraeid gastropod mollusk, Crepidula fornicata, has been widely used for studies of larval developmental biology, physiology, and ecology. Brooded veliger larvae of this species were collected by siphoning onto a sieve after natural release by adults, distributed into the culture at a density of 200/L, and fed with Isochrysis galbana (strain T-ISO) at 1 x 105 cells/mL. Shell growth and acquisition of competence for metamorphosis were documented for sibling larvae reared in ventilated 800 mL cultures designed for equilibration to ambient air or to defined atmospheric gas mixtures. Contrasting with these laboratory culture conditions; growth and competence data were also collected for larvae reared in a 15 L flow-through ambient seawater mesocosm located in a field population of reproductive adults. Growth rates and timing of metamorphic competence in the laboratory cultures were similar to those reported in previously published studies. Larvae reared in the field mesocosm grew much faster and metamorphosed sooner than reported for any laboratory studies. Together, these methods are suited for exploring larval development under predetermined controlled conditions in the laboratory as well as under naturally occurring conditions in the field.

Larval growth and acquisition of competence for metamorphosis were measured in 4 simultaneous replicates of the 800 mL ventilated cultures, each containing 160 larvae, derived from a sibling batch of larvae that hatched from a single egg mass and that were fed Isochrysis galbana at a density of 1 x 105 cells/mL. The pH was 7.9-8.0, temperature was 20-21 &#176;C, and salinity was 30-31 ppt. Growth and metamorphosis were also determined with a different sibling batch of larvae in a single trial of the 1...

Read this Scientific Article Protocol about Laboratory and Field Culture of Larvae of The Slipper Limpet, Crepidula fornicata at JoVE.com

Author Spotlight: Bridging the Gap Between Field Observations and Lab Manipulations in Larval Ecology Research

The paper presents methods for larval culture of the gastropod Crepidula fornicata in a small-volume laboratory-scale system and in an ambient-seawater mesocosm system that can be deployed in the field.

Laboratory and Field Culture of Larvae of The Slipper Limpet, Crepidula fornicata

The calyptraeid gastropod mollusk, Crepidula fornicata, has been widely used for studies of larval developmental biology, physiology, and ecology. Brooded ...

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333 Views.  Dickinson College. The paper presents methods for larval culture of the gastropod Crepidula fornicata in a small-volume laboratory-scale system and in an ambient-seawater mesocosm system that can be deployed in the field.

Video: Author Spotlight: Bridging the Gap Between Field Observations and Lab Manipulations in Larval Ecology Research

High-resolution Measurement of Odor-Driven Behavior in Drosophila Larvae

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral paradigm, the experimenter usually assumes that the rapid diffusion of odorant molecules from the source leads to the creation of a stable gradient in the dish. To establish a quantitative correlation between sensory inputs and behavioral responses, it is necessary to achieve a more thorough characterization of the odorant stimulus conditions. In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

In this video article, we describe a new method allowing the construction of odorant gradients with stable and controllable geometries. We briefly illustrate how these gradients can be used to screen for olfactory defects (full and partial anosmia) and to study more subtle features of chemotaxis behavior.

Olfactory responses in Drosophila larvae have been traditionally studied in Petri dishes comprising a single peripheral odor source. In this behavioral ...

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti. 

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the inner ear (fig 1). Hair cells in the developing cochlea, which are the mechanosensory cells of the auditory system, are aligned in one row of inner hair cells and three (in the base and mid-turns) to four (in the apical turn) rows of outer hair cells that span the length of the organ of Corti. Hair cells transduce sound-induced mechanical vibrations of the basilar membrane into neural impulses that the brain can interpret. Most cases of sensorineural hearing loss are caused by death or dysfunction of cochlear hair cells.
An increasingly essential tool in auditory research is the isolation and in vitro culture of the organ explant 1,2,9. Once isolated, the explants may be utilized in several ways to provide information regarding normative, anomalous, or therapeutic physiology. Gene expression, stereocilia motility, cell and molecular biology, as well as biological approaches for hair cell regeneration are examples of experimental applications of organ of Corti explants.
This protocol describes a method for the isolation and culture of the organ of Corti from neonatal mice. The accompanying video includes stepwise directions for the isolation of the temporal bone from mouse pups, and subsequent isolation of the cochlea, spiral ligament, and organ of Corti. Once isolated, the sensory epithelium can be plated and cultured in vitro in its entirety, or as a further dissected micro-isolate that lacks the spiral limbus and spiral ganglion neurons. Using this method, primary explants can be maintained for 7-10 days. As an example of the utility of this procedure, organ of Corti explants will be electroporated with an exogenous DsRed reporter gene. This method provides an improvement over other published methods because it provides reproducible, unambiguous, and stepwise directions for the isolation, microdissection, and primary culture of the organ of Corti.

Primary Culture and Plasmid Electroporation of the Murine Organ of Corti.

This procedure describes a method for the isolation and culture of the murine organ of Corti with or without the spiral limbus and spiral ganglion neurons. We also demonstrate a method for the expression of an exogenous reporter gene in the organ of Corti explant by electroporation.

In all mammals, the sensory epithelium for audition is located along the spiraling organ of Corti that resides within the conch shaped cochlea of the ...

Microgavage of Zebrafish Larvae

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe interactions17-25. Experimental approaches for studying intestinal biology often require the in vivo introduction of selected materials into the lumen of the intestine. In the larval zebrafish model, this is typically accomplished by immersing fish in a solution of the selected material, or by injection through the abdominal wall. Using the immersion method, it is difficult to accurately monitor or control the route or timing of material delivery to the intestine. For this reason, immersion exposure can cause unintended toxicity and other effects on extraintestinal tissues, limiting the potential range of material amounts that can be delivered into the intestine. Also, the amount of material ingested during immersion exposure can vary significantly between individual larvae26. Although these problems are not encountered during direct injection through the abdominal wall, proper injection is difficult and causes tissue damage which could influence experimental results. We introduce a method for microgavage of zebrafish larvae. The goal of this method is to provide a safe, effective, and consistent way to deliver material directly to the lumen of the anterior intestine in larval zebrafish with controlled timing. Microgavage utilizes standard embryo microinjection and stereomicroscopy equipment common to most laboratories that perform zebrafish research. Once fish are properly positioned in methylcellulose, gavage can be performed quickly at a rate of approximately 7-10 fish/ min, and post-gavage survival approaches 100% depending on the gavaged material. We also show that microgavage can permit loading of the intestinal lumen with high concentrations of materials that are lethal to fish when exposed by immersion. To demonstrate the utility of this method, we present a fluorescent dextran microgavage assay that can be used to quantify transit from the intestinal lumen to extraintestinal spaces. This test can be used to verify proper execution of the microgavage procedure, and also provides a novel zebrafish assay to examine intestinal epithelial barrier integrity under different experimental conditions (e.g. genetic manipulation, drug treatment, or exposure to environmental factors). Furthermore, we show how gavage can be used to evaluate intestinal motility by gavaging fluorescent microspheres and monitoring their subsequent transit. Microgavage can be applied to deliver diverse materials such as live microorganisms, secreted microbial factors/toxins, pharmacological agents, and physiological probes. With these capabilities, the larval zebrafish microgavage method has the potential to enhance a broad range of research fields using the zebrafish model system.

We present a novel method for microgavage of larval zebrafish utilizing standard embryo microinjection and stereomicroscopy equipment. We demonstrate that microgavage is a safe and efficient technique useful for delivering controlled amounts of diverse materials specifically into the larval zebrafish intestinal lumen.

The zebrafish has emerged as a powerful model organism for studying intestinal development1-5, physiology6-11, disease12-16, and host-microbe ...

Design and Development of Aptamer&#8211;Gold Nanoparticle Based Colorimetric Assays for In-the-field Applications

The design and development of an aptamer&#8211;gold nanoparticle (AuNP) colorimetric assay for the detection of small molecules for in-the-field applications was examined. Target selective AuNP based color assays have been developed in controlled proof-of-concept laboratory settings. However, these schemes have not been exerted to a point of failure to determine their practical use beyond laboratory settings. This work describes a generic approach to design, develop, and troubleshoot an aptamer-AuNP colorimetric assay for small molecule analytes and using the assay for in-the-field settings. The assay is advantageous because adsorbed aptamers passivate the nanoparticle surfaces and provide a means to reduce and eliminate false positive responses to non-target analytes. Transitioning this system to practical uses required defining not only the shelf-life of the aptamer-AuNP assay, but establishing methods and procedures for extending the long-term storage capabilities. Also, one of the recognized concerns with colorimetric readout is the burden placed on analysts to accurately identify often subtle changes in color. To lessen the responsibility on analysts in the field, a color analysis protocol was designed to perform the color identification duties without the need for performing this task on laboratory grade equipment. The method for creating and testing the data analysis protocol is described. However to understand and influence the design of adsorbed aptamer assays, the interactions associated with the aptamer, target, and AuNPs require further study. The knowledge gained could lead to tailoring aptamers for improved functionality.

The design and development of an aptamer&#8211;gold nanoparticle colorimetric assay for the detection of small molecules for in-the-field applications was examined. Additionally, a smart-device colorimetric application (app) was validated and long-term storage of the assay was established for use in the field.

The design and development of an aptamer&#8211;gold nanoparticle (AuNP) colorimetric assay for the detection of small molecules for in-the-field ...

Establishment of an Extracellular Acidic pH Culture System

Conditions of the tumor microenvironment, such as hypoxia or nutrient starvation, play critical roles in cancer progression and malignancy. However, the role of acidic extracellular pH in tumor aggressiveness and its underlying mechanism has not been extensively studied compared to hypoxic or nutrient starvation conditions. In addition, a well-defined culture method to mimic the acidic extracellular tumor microenvironment has not been fully reported.
Here we present a simple in vitro culture method to maintain acidic extracellular pH using reduced bicarbonate and increased lactate or HCl concentrations in the culture medium. The medium pH was sustained for at least 24 h and gradually decreased by 72 h following culture of PANC-1 and AsPC-1 pancreatic cancer cells. Three distinct acidic media conditions in this study highly upregulated pH-responsive genes such as MSMO1, INSIG1, and IDI1 compared to hypoxia or nutrient starvation. The upregulation of these genes can be used as a marker of acidic pH. These simple techniques are beneficial to elucidate underlying mechanisms of tumor malignancy under acidic tumor microenvironment. Therefore, our extracellular acidic pH culture system enables discovery of cellular acidic pH responses not only in cancer cells but also in primary cells, such as renal tubular cells, in relation to the other acidic disorders including, diabetic ketoacidosis, lactic acidosis, renal tubular acidosis, and respiratory acidosis.

The acidic tumor microenvironment plays critical roles in tumor progression. To assess the effects of acidic extracellular pH on cancer cells in vitro, we established simple acidic culture systems.

Conditions of the tumor microenvironment, such as hypoxia or nutrient starvation, play critical roles in cancer progression and malignancy. However, the ...

A Protocol for Laboratory Housing of Turquoise Killifish (Nothobranchius furzeri)

The development of husbandry practices in non-model laboratory fish used for experimental purposes has greatly benefited from the establishment of reference fish model systems, such as zebrafish and medaka. In recent years, an emerging fish &#8211; the turquoise killifish (Nothobranchius furzeri) &#8211; has been adopted by a growing number of research groups in the fields of biology of aging and ecology. With a captive life span of 4 - 8 months, this species is the shortest-lived vertebrate raised in captivity and allows the scientific community to test &#8211; in a short time &#8211; experimental interventions that can lead to alterations of the aging rate and life expectancy. Given the unique biology of this species, characterized by embryonic diapause, explosive sexual maturation, marked morphological and behavioral sexual dimorphism - and their relatively short adult life span -&#160;ad hoc husbandry practices are in urgent demand. This protocol reports a set of key husbandry measures that allow optimal turquoise killifish laboratory care, enabling the scientific community to adopt this species as a powerful laboratory animal model.

A Protocol for Laboratory Housing of Turquoise Killifish Nothobranchius furzeri

Laboratory housing of turquoise killifish can be scaled up&#160;to house and efficiently raise thousands of individual fish in a centralized water filtration system, employing the same infrastructure used for standard zebrafish facilities. Here we detail a list of standardized procedures that allow efficient killifish maintenance.

The development of husbandry practices in non-model laboratory fish used for experimental purposes has greatly benefited from the establishment of ...

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and diseases, such as obesity and fatty liver. Usually, lipids are stored in lipid droplets, which are the multifunctional lipid storage organelles in cells. Lipid droplets vary in size and number in different tissues and under different conditions. It has been reported that lipid droplets are tightly controlled through regulation of its biogenesis and degradation. In Drosophila melanogaster, the oenocyte is an important tissue for lipid metabolism and has been recently identified as a human liver analogue regarding lipid mobilization in response to stress. However, the mechanisms underlying the regulation of lipid droplet metabolism in oenocytes remain elusive. To solve this problem, it is of utmost importance to develop a reliable and sensitive method to directly visualize lipid droplet dynamic changes in oenocytes during development and under stressful conditions. Taking advantage of the lipophilic BODIPY 493/503, a lipid droplet-specific fluorescent dye, described here is a detailed protocol for the dissection and subsequent lipid droplet staining in the oenocytes of Drosophila larvae in response to starvation. This allows for qualitative analysis of lipid droplet dynamics under various conditions by confocal microscopy. Furthermore, this rapid and highly reproducible method can also be used in genetic screens for indentifying novel genetic factors involving lipid droplet metabolism in oenocytes and other tissues.

Dissection and Lipid Droplet Staining of Oenocytes in Drosophila Larvae

Presented here are detailed methods for the dissection and lipid droplet staining of oenocytes in Drosophila larvae using BODIPY 493/503, a lipid droplet-specific fluorescent dye.

Lipids are essential for animal development and physiological homeostasis. Dysregulation of lipid metabolism results in various developmental defects and ...

Development of a Mobile Mitochondrial Physiology Laboratory for Measuring Mitochondrial Energetics in the Field

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to investigate metabolic capability. To obtain the measures of mitochondrial respiration, fresh biological samples must be used, and the entire laboratory procedure must be completed within approximately 2 h. Furthermore, multiple pieces of specialized equipment are required to perform these laboratory assays. This creates a challenge for measuring mitochondrial respiration in the tissues of wild animals living far from physiology laboratories as live tissue cannot be preserved for very long after collection in the field. Moreover, transporting live animals over long distances induces stress, which can alter mitochondrial energetics.
This manuscript introduces the Auburn University (AU) MitoMobile, a mobile mitochondrial physiology laboratory that can be taken into the field and used on-site to measure mitochondrial metabolism in tissues collected from wild animals. The basic features of the mobile laboratory and the step-by-step methods for measuring isolated mitochondrial respiration rates are presented. Additionally, the data presented validate the success of outfitting the mobile mitochondrial physiology laboratory and making mitochondrial respiration measurements. The novelty of the mobile laboratory lies in the ability to drive to the field and perform mitochondrial measurements on the tissues of animals captured on site.

We designed and constructed a mobile laboratory to measure respiration rates in isolated mitochondria of wild animals captured at field locations. Here, we describe the design and outfitting of a mobile mitochondrial laboratory and the associated laboratory protocols.

Mitochondrial energetics is a central theme in animal biochemistry and physiology, with researchers using mitochondrial respiration as a metric to ...

Cell Culture Techniques and Practices to Avoid Contamination by Fungi and Bacteria in the Research Cell Culture Laboratory

Cell culture is a delicate skill necessary for growing human, animal, and insect cells, or other tissues, in a controlled environment. The goal of the protocol is to emphasize the correct techniques used in a research laboratory to prevent contamination from fungi and bacteria. Special emphasis is placed on avoiding mycoplasma contamination, a major concern in the cell culture room due to its small size and resistance to most antibiotics used for cell culture. These same techniques ensure continuous growth and maintain healthy cells. For new and experienced cell culture users alike, it&#8217;s important to consistently adhere to these best practices to mitigate the risk of contamination. Once a year, laboratories should review cell culture best practices and follow-up with a discussion or additional training if needed. Taking early action to prevent contamination in the first place will save time and money, as compared to cleaning up after contamination occurs. Universal best practices keep cell cultures healthy, thereby reducing the need to constantly thaw new cells, purchase expensive cell culture media, and reducing the amount of incubator decontamination and downtime.&#160;

This protocol presents essential cell culture techniques and practices to be used in the research cell culture laboratory to avoid contamination by fungi and bacteria. Within the category of bacteria, special emphasis will be placed on preventing mycoplasma contamination.&#160;&#160;

Cell culture is a delicate skill necessary for growing human, animal, and insect cells, or other tissues, in a controlled environment. The goal of the ...

The Mushing, Sieving, and Floating Method for Detecting Fruit Fly Larvae

Sieving Fruit Pulp to Detect Immature Tephritid Fruit Flies in the Field

Fruit flies of the Tephritidae family are among the most destructive and invasive agricultural pests in the world. Many countries undertake expensive eradication programs to eliminate incipient populations. During eradication programs, a concerted effort is made to detect larvae, as this strongly indicates a breeding population and helps establish the spatial extent of the infestation. The detection of immature life stages triggers additional control and regulatory actions to contain and prevent any further spread of the pest. Traditionally, larval detection is accomplished by cutting individual host fruits and examining them visually. This method is labor intensive, as only a limited number of fruit can be processed, and the probability of missing a larva is high. An extraction technique that combines i) mushing host fruit in a plastic bag, ii) straining pulp through a series of sieves, iii) placing retained pulp in a brown sugar water solution, and iv) collecting larvae that float to the surface was tested. The method was evaluated in Florida with field-collected guava naturally infested by Anastrepha suspensa. To mimic low populations more representative of a fruit fly eradication program, mangos and papaya in Hawaii were infested with a known, low number of Bactrocera dorsalis larvae. The applicability of the method was tested in the field on guava naturally infested by B. dorsalis to evaluate the method under conditions experienced by workers during an emergency fruit fly program. In both field and laboratory trials, mushing and sieving the pulp was more efficient (required less time) and more sensitive (more larvae found) than cutting fruit. Floating the pulp in brown sugar water solution helped detect earlier instar larvae. Mushing and sieving fruit pulp of important tephritid hosts may increase the probability of detecting larvae during emergency programs.

Author Spotlight: Sieving Fruit Pulp to Detect Immature Tephritid Fruit Flies in the Field  

Increasing the detection of immature tephritid fruit flies in the field can trigger timely efforts to eliminate populations of these destructive pests. Detecting late instar larvae is faster and more accurate when mushing host fruit in a bag and passing the pulp through a series of sieves than hand cutting and visual inspection.

Fruit flies of the Tephritidae family are among the most destructive and invasive agricultural pests in the world. Many countries undertake expensive ...