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

Summary

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.

Abstract

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.

Introduction

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 development^2,3. 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 America^4,5. 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 performance^{6,7,8,9,10,11}. 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 animal^{12,13,14,15,16}.

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.

Protocol

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.

1. Make a sieve by cutting the bottom off of a 400 mL tri-corner disposable plastic beaker and gluing on a panel of 236 µm nylon mesh. For this purpose and especially for mesocosm construction (below), use a hot melt glue that is formulated for good adhesion to polyethylene.
2. 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.
3. Siphon the larvae through a short length (15-20 cm) of 5 mm glass tubing affixed to the end of a 60-80 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.
4. 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 °C^17,18. Filtration at < 1 µm is sufficient to eliminate fouling epizootic protists that can cause problems in laboratory cultures.
5. 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.
6. 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.
7. Change culture water every other day by pouring the contents of the culture into a sieve within a beaker, as in step 1.3 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.
   ​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.

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.

1. Drill two 5 mm holes in the lid of each culture jar (including the lid'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's drill bit will also make an appropriate clearance hole).
2. 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.
3. 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.
4. 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.
5. 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.
6. 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.

3. Construction of a field-deployable mesocosm culture for larvae of C. fornicata

1. 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.
2. Cut out 4 panels of 236 µm 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.
3. 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.
4. 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.
5. 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.
6. 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.
7. 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 2/3^rd out of the water and repeatedly bailing from its bottom into the fresh mesocosm with a 2 L pitcher.
8. 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.

Results

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 °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...

Discussion

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 essential^17,19. 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...

Materials

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