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/pubmed?term=((The+embryology+of+Crepidula%3A+a+contribution+to+the+cell+lineage+and+early+development+of+some+marine+gastropods%5BTitle%5D)+AND+%22Gasteropods%22%5BJournal%5D)">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/pubmed?term=((The+slipper+snail%2C+Crepidula%3A+an+emerging+lophotrochozoan+model+system%5BTitle%5D)+AND+%22Biol+Bull%22%5BJournal%5D)">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/pubmed?term=((Slipper+snail+tales%3A+how+Crepidula+fornicata+and+Crepidula+atrasolea+became+model+molluscs%5BTitle%5D)+AND+%22Curr+Top+Dev+Biol%22%5BJournal%5D)">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/pubmed?term=((Spread+of+the+slipper+limpet+Crepidula+fornicata+%28L.+1758%29+in+Europe.+Current+state+and+consequences%5BTitle%5D)+AND+%22ScientiaMarina%22%5BJournal%5D)">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/pubmed?term=((Field+reproductive+dynamics+of+the+invasive+slipper+limpet%2C+Crepidula+fornicata%5BTitle%5D)+AND+%22J+Exp+Mar+Biol+Ecol%22%5BJournal%5D)">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/pubmed?term=((The+relationship+between+temperature%2C+growth+rate%2C+and+duration+of+planktonic+life+for+larvae+of+the+gastropod+Crepidula+fornicata+%28L.%29%5BTitle%5D)+AND+%22J+Exp+Mar+Biol+Ecol%22%5BJournal%5D)">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/pubmed?term=((Latent+effects%3A+surprising+consequences+of+embryonic+and+larval+experience+on+life+after+metamorphosis%5BTitle%5D)+AND+%22Evol+Ecol+Marine+Invertebrate+Larvae%22%5BJournal%5D)">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/pubmed?term=((Onset+of+metamorphic+competence+in+larvae+of+the+gastropod+Crepidula+fornicata+%28L.%29%2C+judged+by+a+natural+and+an+artificial+cue%5BTitle%5D)+AND+%22J+Exp+Mar+Biol+Ecol%22%5BJournal%5D)">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/pubmed?term=((Inhibitory+function+of+nitric+oxide+on+the+onset+of+metamorphosis+in+competent+larvae+of+Crepidula+fornicata%3A+A+transcriptional+perspective%5BTitle%5D)+AND+%22Mar+Genomics%22%5BJournal%5D)">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/pubmed?term=((Neural+correlates+of+settlement+in+veliger+larvae+of+the+gastropod%2C+Crepidula+fornicata%5BTitle%5D)+AND+%22Invertebrate+Biol%22%5BJournal%5D)">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/pubmed?term=((Carryover+effects+of+brooding+conditions+on+larvae+in+the+slipper+limpet+Crepidula+fornicata%5BTitle%5D)+AND+%22Mar+Ecol+Prog+Ser%22%5BJournal%5D)">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/pubmed?term=((Does+encapsulation+protect+embryos+from+the+effects+of+ocean+acidification%3F+The+example+of+Crepidula+fornicata%5BTitle%5D)+AND+%22PLoS+One%22%5BJournal%5D)">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/pubmed?term=((Resilience+of+Atlantic+slippersnail+Crepidula+fornicata+larvae+in+the+face+of+severe+coastal+acidification%5BTitle%5D)+AND+%22Front+in+Mar+Sci%22%5BJournal%5D)">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/pubmed?term=((Legacy+of+multiple+stressors%3A+Responses+of+gastropod+larvae+and+juveniles+to+ocean+acidification+and+nutrition%5BTitle%5D)+AND+%22Biol+Bull%22%5BJournal%5D)">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/pubmed?term=((Impact+of+ocean+acidification+on+growth%2C+onset+of+competence%2C+and+perception+of+cues+for+metamorphosis+in+larvae+of+the+slippershell+snail%2C+Crepidula+fornicata%5BTitle%5D)+AND+%22Mar+Biol%22%5BJournal%5D)">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/pubmed?term=((The+marine+gastropod+Crepidula+fornicata+remains+resilient+to+ocean+acidification+across+two+life+history+stages%5BTitle%5D)+AND+%22Front+Physiol%22%5BJournal%5D)">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/pubmed?term=((Artificial+seawater+culture+of+the+gastropod+Crepidula+fornicata+for+studies+of+larval+settlement+and+metamorphosis.+In%3A+Carroll%2C+D.%2C+Stricker%2C+S.+%28eds%29+Developmental+biology+of+the+sea+urchin+and+other+marine+invertebrates%5BTitle%5D)+AND+%22Methods+Mol+Biol%22%5BJournal%5D)">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/pubmed?term=((The+interactive+influence+of+temperature+and+salinity+on+larval+and+juvenile+growth+in+the+gastropod+Crepidula+fornicata+%28L.%29%5BTitle%5D)+AND+%22J+Exp+Mar+Biol+Ecol%22%5BJournal%5D)">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://scholar.google.com/scholar?hl=en&safe=off&q=author%3aStrathmann%2C+MF+%22Reproduction+and+Development+of+Marine+Invertebrates+of+the+Northern+Pacific+Coast%3A+Data+and+Methods+for+the+Study+of+Eggs%2C+Embryos%2C+and+Larvae%22">Reproduction and Development of Marine Invertebrates of the Northern Pacific Coast: Data and Methods for the Study of Eggs, Embryos, and Larvae</a>. , University of Washington Press. (1987).</li>
<li>Clark, H. R., Gobler, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Diurnal+fluctuations+in+CO2+and+dissolved+oxygen+concentrations+do+not+provide+a+refuge+from+hypoxia+and+acidification+for+early-life-stage+bivalves%5BTitle%5D)+AND+%22Mar+Ecol+Prog+Ser%22%5BJournal%5D)">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/pubmed?term=((Hypoxia+and+acidification+in+ocean+ecosystems%3A+coupled+dynamics+and+effects+on+marine+life%5BTitle%5D)+AND+%22Biol+Lett%22%5BJournal%5D)">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/pubmed?term=((Relationship+between+larval+and+juvenile+growth+rates+in+two+marine+gastropods%2C+Crepidula+plana+and+C%5BTitle%5D)+AND+%22fornicata.+Mar+Biol%22%5BJournal%5D)">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/pubmed?term=((Effect+of+food+on+metamorphic+competence+in+the+model+system+Crepidula+fornicata%5BTitle%5D)+AND+%22Biol+Bull%22%5BJournal%5D)">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/pubmed?term=((Coastal+ocean+acidification%3A+the+other+eutrophication+problem%5BTitle%5D)+AND+%22Estuarine%2C+Coast+Shelf+Sci%22%5BJournal%5D)">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, bright, and free of visible fouling by stalked ciliate protists or sessile pennate diatoms, which spread rapidly in crowded cultures. Shell fouling inhibits swimming and feeding, causes mortality, and is usually an indication that the density of larvae in the culture is too high.
The larval growth rates and timing of competence documented here in the ventilated laboratory cultures are similar to published results from laboratory experiments using an earlier version of this method under similar conditions of larval density, pH, temperature, salinity, and food ration13,14,16. The only substantive difference between the method described here and the one used in the latter studies is that in the present experiment, the ventilating gas stream was introduced through a 2 mm (outer diameter) tube that created a stream of bubbles from the bottom of the culture jar, rather than being passed across the headspace at the surface of the jar. The water circulation and mixing created by the bubble stream is thus not deleterious to larval growth and acquisition of competence and may have advantages for experiments requiring rapid equilibration of culture seawater to changing gas mixes, e.g., for experiments investigating effects of diel cycles in the partial pressures of CO2 and dissolved O2 such as occur in productive nearshore marine environments20,21.
Mesocosm results yielded larval growth rates in excess of any that have been published from laboratory studies, as well as the shortest time to acquisition of competence for metamorphosis22,23. Although the present results were obtained under field conditions that were clearly very favorable for larval growth and development, the method should be most informative for exploring larval performance in field sites that exhibit naturally-occurring combinations of environmental stressors, including at-risk and degraded habitats that are of interest for purposes of management and remediation24.

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><tbody><tr><td>Bucket, Polyethylene, 7 gallon</td><td>US Plastic</td><td>16916</td><td>for mesocosm</td></tr><tr><td>&lt;em&gt;Crepidula fornicata&lt;/em&gt;</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 &amp;micro;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&quot; diameter, 5/32&quot;-5/16&quot; 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&quot; 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&quot;x3/16&quot;x1/32&quot;</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 15 L mesocosm containing 600 larvae, deployed in Buzzards Bay, MA, USA, under similar conditions of pH, temperature, and salinity as the lab cultures, but fed by natural phytoplankton in the ambient seawater that flowed through the mesh panels of the mesocosm. The pH measured was 7.8-8.3, temperature was 20-24 &#176;C, and salinity was 26-28 ppt. Growth was determined as a change in shell length (Figure 3). Larvae grew faster in the mesocosm (71 &#181;m/day) than in the laboratory cultures (54 &#181;m/day) during the first 4-5 days after hatching (Figure 4). Between the 5th and 6th days, the larvae in the mesocosm began to metamorphose spontaneously, and the larvae that remained in the mesocosm on day 6 were, on average, 181 &#181;m bigger than on day 5. There were no further larval measurements from the mesocosm because most individuals had metamorphosed by day 7. Larvae in the laboratory cultures grew at a much slower rate from day 4 to day 8 (41 &#181;m/day) and from day 8 to day 13 (31 &#181;m/day). These larvae began to become competent for metamorphosis on day 8 and were nearly all competent by day 12, as determined by stimulation of subsamples with elevated [K+], described in8. There was little spontaneous metamorphosis in the laboratory cultures (&#60;10%) through day 13, when the experiment was terminated. Survivorship was 91%-100% in the lab cultures and was not determined in the mesocosm because of the difficulty in inspecting and recovering all individuals from the mesocosm volume and surfaces.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66208/66208fig01.jpg" /> 
Figure 1: Ventilated culture setup. The figure shows 800 mL ventilated cultures containing larvae of Crepidula fornicata, stocked at a density of 1 larva/5 mL. Note a thin stream of bubbles, especially visible in jars 2 and 4. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66208/66208fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66208/66208fig02.jpg" /> 
Figure 2: Field-deployable mesocosm culture setup. The figure shows 15 L mesocosm, (A) lateral view, (B) top view, and (C) setup deployed in floating rack. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66208/66208fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66208/66208fig03.jpg" /> 
Figure 3: Veliger larva of Crepidula fornicata at 4 days post-hatching. Dashed line indicates axis of shell length measurement. Abbreviations: s = shell; v = velum; f = foot; o = operculum. Scale bar = 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66208/66208fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66208/66208fig04.jpg" /> 
Figure 4: Growth of larvae of Crepidula fornicata in lab and field cultures. Growth of larvae in&#160;800 mL laboratory cultures (dashed line, open circles) and 15 L field mesocosm (solid line, filled squares). Each point represents the mean of 15 larvae from each of 4 replicate laboratory cultures or the mean of 25 larvae from a single mesocosm. Error bars are &#177; 1 SD. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/66208/66208fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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 ...

laboratory-field-culture-larvae-slipper-limpet-crepidula

Nanyang Technological University

Research

JoVE Journal

Biology

376 Views.  Dickinson College. Ik ben geïnteresseerd in hoe de ontwikkeling van mariene larven wordt beïnvloed door meerdere omgevingsfactoren en stressoren en hoe het larvale zenuwstelsel informatie uit de omgeving integreert om de vestiging van larven, metamorfose naar benthische juvenielen en rekrutering in volwassen populaties te reguleren. Larven van sommige mariene soorten zoals de pantoffellimpet, Crepidula fornicata, zijn vrij resistent tegen verzuring van de oceaan, maar er zijn belangrijke interacties tussen ve...

Video: Auteur in de schijnwerpers: de kloof overbruggen tussen veldobservaties en laboratoriummanipulaties in onderzoek naar larvale ecologie

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. This is due to its reproductive behavior: females damage the external surface of fruits and vegetables when they lay eggs and the hatched larvae feed on their pulp. Wild C. capitata populations are traditionally controlled through insecticide spraying and/or eco-friendly approaches, the most successful being the Sterile Insect Technique (SIT). The SIT relies on mass-rearing, radiation-based sterilization and field release of males that retain their capacity to mate but are not able to generate fertile progeny. The advent and the subsequent rapid development of biotechnological tools, together with the availability of the medfly genome sequence, has greatly boosted our understanding of the biology of this species. This favored the proliferation of new strategies for genome manipulation, which can be applied to population control.
In this context, embryo microinjection plays a dual role in expanding the toolbox for medfly control. The ability to interfere with the function of genes that regulate key biological processes, indeed, expands our understanding of the molecular machinery underlying medfly invasiveness. Furthermore, the ability to achieve germ-line transformation facilitates the production of multiple transgenic strains that can be tested for future field applications in novel SIT settings. Indeed, genetic manipulation can be used to confer desirable traits that can, for example, be used to monitor sterile male performance in the field, or that can result in early life-stage lethality. Here we describe a method to microinject nucleic acids into medfly embryos to achieve these two main goals.

Delivery of Nucleic Acids through Embryo Microinjection in the Worldwide Agricultural Pest Insect, Ceratitis capitata

The Mediterranean fruit fly (medfly) Ceratitis capitata (Diptera: Tephritidae) is a worldwide pest of agriculture. A deeper understanding of its biology is key to control medfly populations and thus reduce economic impact. Embryo microinjection is a fundamental tool allowing both germ-line transformation and reverse genetics studies in this species.

The Mediterranean fruit fly (medfly) Ceratitis capitata (Wiedemann) (Diptera: Tephritidae) is a pest species with extremely high agricultural relevance. ...

Genetics

Use of Chironomidae (Diptera) Surface-Floating Pupal Exuviae as a Rapid Bioassessment Protocol for Water Bodies

Rapid bioassessment protocols using benthic macroinvertebrate assemblages have been successfully used to assess human impacts on water quality. Unfortunately, traditional benthic larval sampling methods, such as the dip-net, can be time-consuming and expensive. An alternative protocol involves collection of Chironomidae surface-floating pupal exuviae (SFPE). Chironomidae is a species-rich family of flies (Diptera) whose immature stages typically occur in aquatic habitats. Adult chironomids emerge from the water, leaving their pupal skins, or exuviae, floating on the water&#8217;s surface. Exuviae often accumulate along banks or behind obstructions by action of the wind or water current, where they can be collected to assess chironomid diversity and richness. Chironomids can be used as important biological indicators, since some species are more tolerant to pollution than others. Therefore, the relative abundance and species composition of collected SFPE reflect changes in water quality. Here, methods associated with field collection, laboratory processing, slide mounting, and identification of chironomid SFPE are described in detail. Advantages of the SFPE method include minimal disturbance at a sampling area, efficient and economical sample collection and laboratory processing, ease of identification, applicability in nearly all aquatic environments, and a potentially more sensitive measure of ecosystem stress. Limitations include the inability to determine larval microhabitat use and inability to identify pupal exuviae to species if they have not been associated with adult males.

Use of Chironomidae Diptera Surface-Floating Pupal Exuviae as a Rapid Bioassessment Protocol for Water Bodies

Rapid bioassessment protocols using benthic macroinvertebrates are often used to monitor and assess water quality. An efficient protocol involves collections of Chironomidae surface-floating pupal exuviae (SFPE). Here, techniques for field collection, laboratory processing, slide mounting, and identification of Chironomidae SFPE are described.

Rapid bioassessment protocols using benthic macroinvertebrate assemblages have been successfully used to assess human impacts on water quality. ...

Environment

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

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

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

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

Copepods of the genus Tigriopus, which are common zooplankton in rocky tide pools, show precopulatory mate-guarding behavior where a male clasps a ...

Use of Galleria mellonella as a Model Organism to Study Legionella pneumophila Infection

Legionella pneumophila, the causative agent of a severe pneumonia named Legionnaires&#39; disease, is an important human pathogen that infects and replicates within alveolar macrophages. Its virulence depends on the Dot/Icm type IV secretion system (T4SS), which is essential to establish a replication permissive vacuole known as the Legionella containing vacuole (LCV). L. pneumophila infection can be modeled in mice however most mouse strains are not permissive, leading to the search for novel infection models. We have recently shown that the larvae of the wax moth Galleria mellonella are suitable for investigation of L. pneumophila infection. G. mellonella is increasingly used as an infection model for human pathogens and a good correlation exists between virulence of several bacterial species in the insect and in mammalian models. A key component of the larvae&#39;s immune defenses are hemocytes, professional phagocytes, which take up and destroy invaders. L. pneumophila is able to infect, form a LCV and replicate within these cells. Here we demonstrate protocols for analyzing L. pneumophila virulence in the G. mellonella model, including how to grow infectious L. pneumophila, pretreat the larvae with inhibitors, infect the larvae and how to extract infected cells for quantification and immunofluorescence microscopy. We also describe how to quantify bacterial replication and fitness in competition assays. These approaches allow for the rapid screening of mutants to determine factors important in L. pneumophila virulence, describing a new tool to aid our understanding of this complex pathogen.

Use of Galleria mellonella as a Model Organism to Study Legionella pneumophila Infection

The larva of the wax moth Galleria mellonella was recently established as an in vivo model to study Legionella pneumophila infection. Here, we demonstrate fundamental techniques to characterize the pathogenesis of Legionella in the larvae, including inoculation, measurement of bacterial virulence and replication as well as extraction and analysis of infected hemocytes.

Legionella pneumophila, the causative agent of a severe pneumonia named Legionnaires&#39; disease, is an important human pathogen that infects and ...

Immunology and Infection

Exploring Life History Choices: Using Temperature and Substrate Type as Interacting Factors for Blowfly Larval and Female Preferences

Blowflies (Diptera: Calliphoridae) present a wide range of larval lifestyles, typically classified as obligate parasitism, facultative parasitism, and complete sapro-necrophagy. Several parasitic species, both obligate and facultative, are considered to be of sanitary and economic importance, as their larvae can cause myiasis (maggot infestation in live tissue). However, it is noteworthy that the adult female plays a decisive role as she chooses the oviposition site, and, therefore, largely determines the feeding habit and developmental conditions of the larvae. In this study, two protocols are proposed to test larval feeding preference and female oviposition site preference considering two interacting factors: meat substrate type and temperature. The setups presented here allowed to test Lucilia cuprina larvae and gravid females in a four-choice assay with two temperatures (33 &plusmn; 2 &#176;C and 25 &plusmn; 2 &#176;C) and two types of meat substrates (fresh meat supplemented with blood and 5-day-old rotten meat). Larvae or gravid females can choose to burrow or lay their eggs, respectively, in either of the following: rotten meat at 25 &#176;C (simulating a necrophagous species condition), fresh meat supplemented with blood at 33 &#176;C (simulating a parasitic species condition), and two controls, rotten meat at 33 &#176;C, or fresh meat supplemented with blood at 25 &#176;C. The preference is assessed by counting the number of larvae or eggs laid in each option for each replicate. Comparing the observed results to a random distribution allowed for the estimation of the statistical significance of the preference. The results indicated that L. cuprina larvae have a strong preference for the rotten substrate at 25 &#176;C. Conversely, oviposition-site preference by females was more varied for the meat type. This methodology can be adapted to test the preference of other insect species of similar size. Other questions can also be explored by using alternative conditions.

Herein, two protocols for assessing food source and oviposition preferences in larvae and females of blowflies are detailed. These comprise four choices with two interacting factors: substrate type and temperature. The assays enable the determination of the food source preference of the larvae and the oviposition site preference for the females.

Blowflies (Diptera: Calliphoridae) present a wide range of larval lifestyles, typically classified as obligate parasitism, facultative parasitism, and ...

Behavior

The Galleria mellonella Waxworm Infection Model for Disseminated Candidiasis

Candida species are common fungal commensals of humans colonizing the skin, mucosal surfaces, and gastrointestinal tract. Under certain conditions, Candida can overgrow their natural niches resulting in debilitating mucosal infections as well as life-threatening systemic infections, which are a major focus of investigation due to their associated high mortality rates. Animal models of disseminated infection exist for studying disease progression and dissecting the characteristics of Candida pathogenicity. Of these, the Galleria mellonella waxworm infection model provides a cost-effective experimental tool for high-throughput investigations of systemic virulence. Many other bacterial and eukaryotic infectious agents have been effectively studied in G. mellonella to understand pathogenicity, making it a widely accepted model system. Yet, variation in the method used to infect G. mellonella can alter phenotypic outcomes and complicate interpretation of the results. Here, we outline the benefits and drawbacks of the waxworm model to study systemic Candida pathogenesis and detail an approach to improve reproducibility. Our results highlight the range of mortality kinetics in G. mellonella and describe the variables which can modulate these kinetics. Ultimately, this method stands as an ethical, rapid, and cost-effective approach to study virulence in a model of disseminated candidiasis.

The Galleria mellonella Waxworm Infection Model for Disseminated Candidiasis

Galleria mellonella serves as an invertebrate model for disseminated candidiasis. Here, we detail the infection protocol and provide supporting data for the model&#39;s effectiveness.

Candida species are common fungal commensals of humans colonizing the skin, mucosal surfaces, and gastrointestinal tract. Under certain conditions, ...

Laboratory Rearing of Stable Flies and Other Muscoid Diptera

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in agronomic practices resulted in serious outbreaks of stable flies in several countries. These outbreaks disrupted livestock production and human recreation resulting in public demands for increasing research and management efforts for this pest. A simple and inexpensive procedure for rearing stable flies for laboratory studies is presented. The procedure uses locally available diet components, equipment and supplies. The procedure can be adapted for rearing other muscoid flies including face fly (Musca autumnalis), horn fly (Haematobia irritans), and house fly (Musca domestica). The procedure produces stable fly puparia averaging 12.5 mg and ~35% egg to adult survival. Approximately 3000 flies are produced in each pan.

A procedure for rearing stable flies (Stomoxys calcitrans) is presented. The procedure uses locally available materials for diet components, equipment and supplies.

Stable flies, Stomoxys calcitrans, are serious pests of livestock, humans, companion animals and wildlife worldwide. During the last 20+ years, changes in ...

At-Risk Butterfly Captive Propagation Programs to Enhance Life History Knowledge and Effective Ex Situ Conservation Techniques

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. Research on such captive populations can also yield valuable data to address key information gaps about the behavior, life history, and ecology of the target taxa. We describe a protocol for captive propagation of the federally endangered Cyclargus thomasi bethunebakeri that can be used as a model for other at-risk butterfly ex situ programs, especially those in the family Lycaenidae. We further provide a simple and straightforward protocol for recording various life history metrics that can be useful for informing ex situ methodologies as well as adapted for laboratory studies of other lepidoptera.

Here, we present protocols for 1) the laboratory captive propagation of the federally endangered Miami blue butterfly (Cyclargus thomasi bethunebakeri), and 2) assessing basic life history information such as immature development time and number of larval stadia. Both methods can be adapted for use with other ex situ conservation programs.

Improving knowledge of ex situ best practices for at-risk butterflies is important for generating successful conservation and recovery program outcomes. ...

Preparing and Rearing Axenic Insects with Tissue Cultured Seedlings for Host-Gut Microbiota Interaction Studies of the Leaf Beetle

Insect guts are colonized by diverse bacteria that can profoundly impact the host&#39;s physiological traits. Introducing a particular bacterial strain into an axenic insect is a powerful method to verify gut microbial function and elucidate the mechanisms underlying gut microbe-host interactions. Administering antibiotics or sterilizing egg surfaces are two commonly used methods to remove gut bacteria from insects. However, in addition to the potential adverse effects of antibiotics on insects, previous studies indicated that feeding antibiotics could not eliminate gut bacteria. Thus, germ-free artificial diets are generally employed to maintain axenic insects, which is a tedious and labor-intensive process that cannot fully resemble nutritional components in natural food. Described here is an efficient and simple protocol to prepare and maintain axenic larvae of a leaf beetle&#160;(Plagiodera versicolora). Specifically, surfaces of the beetle eggs were sterilized, following which germ-free poplar leaves were used to rear axenic larvae. The axenic status of the insects was further confirmed via culture-dependent and culture-independent assays. Collectively, by combining egg disinfection and germ-free cultivation, an efficient and convenient method was developed to obtain axenic P. versicolora, providing a readily transferable tool for other leaf-eating insects.

To obtain an axenic insect, its egg surface is sterilized, and the hatched larva is subsequently reared using axenic leaves. This method provides an efficient way for axenic insect preparation without administering antibiotics or developing an artificial diet, which can also be applied to other leaf-eating insects.

Insect guts are colonized by diverse bacteria that can profoundly impact the host&#39;s physiological traits. Introducing a particular bacterial strain ...

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human food. Outside its native range, culturing G. bimaculatus is feasible due to its dietary plasticity, rapid reproductive cycle, lack of diapause requirement, tolerance for high-density rearing, and robustness against pathogens. Thus, G. bimaculatus can be&#160;a versatile model for studies of insect physiology, behavior, embryology, or genetics.
Cultural parameters, such as stocking density, within-cage refugia, photoperiod, temperature, relative humidity, and diet, all impact cricket growth, behavior, and gene expression and should be standardized. In the burgeoning literature on farming insects for human consumption, these crickets are frequently employed to evaluate candidate feed admixtures derived from crop residues, food-processing byproducts, and other low-cost waste streams.
To support ongoing experiments evaluating G. bimaculatus growth performance and nutritional quality in response to variable feed substrates, a comprehensive set of standard protocols for breeding, upkeep, handling, measurement, and euthanasia in the laboratory was developed and is presented here. An industry-standard cricket feed has proven nutritionally adequate and functionally appropriate for the long-term maintenance of cricket breeding stocks, as well as for use as an experimental control feed. Rearing these crickets at a density of 0.005 crickets/cm3 in screen-topped 29.3 L polyethylene cages at an average temperature of 27 &#176;C on a 12 light (L)/12 dark (D) photoperiod, with moistened coconut coir serving both as hydration source and oviposition medium has successfully sustained healthy crickets over a 2-year span. Following these methods, crickets in a controlled experiment yielded an average mass of 0.724 g 0.190 g at harvest, with 89% survivorship and 68.2% sexual maturation between stocking (22 days) and harvest (65 days).

Maintaining Laboratory Cultures of Gryllus bimaculatus, a Versatile Orthopteran Model for Insect Agriculture and Invertebrate Physiology

This paper outlines basic methods to standardize important factors such as density, feed availability, hydration source, and environmental controls for the long-term rearing of laboratory cultures of the edible cricket, Gryllus bimaculatus.

Gryllus bimaculatus (De Geer)&#160;is a large-bodied cricket distributed throughout Africa and Southern Eurasia where it is often wild-harvested as human ...

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

Samenvatting

Meer video's verkennen