|California State University Long Beach (CSULB)
|Principal Investigator, Contact
|California State University Long Beach (CSULB)
|Woods Hole Oceanographic Institution (WHOI BCO-DMO)
|BCO-DMO Data Manager
Collection of adults and spawning: Adults of each species were collected from intertidal or shallow subtidal zones from various sites in Los Angeles County and transported to California State University Long Beach, where they were maintained in recirculating seawater tanks at 16 °C until their use in experiments. Experiments were carried out on one species at a time, depending on reproductive seasonality for that species.
Spawning was induced using standard methods (e.g., M. Strathmann, 1987). The echinoids D**endraster excentricus, Lytechnius pictus, Strongylocentrotus purpuratus and S. fragilis were induced to spawn via injection of 0.2-1.0 mL (depending on adult size) 0.53 M KCL into the perivisceral coelom. The asteroids Patiria miniata and Astropecten armatus were induced to spawn by injection of 1-3 mL 100 µM 1-methyladenine. The holothuroid Apostichopus parvimensis was injected with 3 mL of 200 µM NGLWY-amide (Kato et al., 2009). The ophiuroid Ophiothrix spiculata was exposed to 4 °C water in the dark for 15 minutes, then to room temperature water and sunlight for 15 minutes; this treatment was repeated for up to two hours (Selvakumaraswamy & Byrne, 2000). For all species, adults were each induced to spawn in their own isolated containers, allowing us to control subsequent fertilizations.
Quantification of ciliated band length per unit protein
Design: All eight species listed above were studied with the aim of comparing the length of ciliated band supported per unit protein between larval forms (pluteus and non-pluteus). For each species, we prepared a genetically diverse population from at minimum three parents of each sex. Eggs of each female were rinsed in 0.2 µm filtered seawater (FSW) and resuspended separately in a small volume of FSW. Three drops of concentrated sperm were collected from each male and combined together in 50 mL FSW. The sperm mixture was then added in small increments to each set of eggs until a fertilization rate of ~90% was reached. The fertilized eggs were then diluted in FSW and placed in an environmental chamber at 16 °C.
Twenty-four hours after fertilization, swimming embryos were decanted into clean beakers to separate them from unfertilized gametes or dead embryos. The beakers were then stirred to evenly distribute the embryos, and the concentration of embryos in each beaker was estimated based on counts in five 0.5 mL aliquots. Equal numbers of embryos from each mother were then added to each of seven replicate 2 L beakers for a total of 500 embryos of mixed parentage per beaker. For S. purpuratus, 14 replicate 2 L beakers were generated since larvae were small and more were needed for protein analysis. Embryos were fed 6000 cells mL−1 of Rhodomonas lens grown under natural light in F/2 medium. Algal cells were separated from the culture medium by centrifugation then resuspended in FSW before their concentration was estimated using a flow cytometer (BD Accuri 6). All beakers were then maintained in an environmental chamber at 16 °C, with each beaker being stirred by a paddle system operating at ~4 strokes per minute (M. Strathmann, 1987).
D**aily maintenance:** After the initial setup of cultures, water changes and reestablishment of food treatments occurred daily beginning 3 days post fertilization (dpf). The contents of each beaker were poured into a submerged 60 µm sieve to concentrate larvae, which were then gently rinsed back into the cleaned beaker with fresh FSW. Cultures were then fed 6000 cells mL−1 of Rhodomonas lens as above and returned to the paddle system.
Protein analysis: For all species except D. excentricus, sampling for images and protein analysis occurred at 5 and 10 dpf during the water change while larvae were concentrated. Sampling for D. excentricus occurred at 3 and 5 dpf due to their more rapid development and earlier formation of the juvenile structures. We collected larvae using a pulled pipette while observing them with a dissection microscope. Depending on larval size and age, three samples of 30-80 larvae each were collected from each beaker and placed in 1.5 mL microcentrifuge tubes on ice. Exact counts of larvae for each tube were noted. The remaining larvae were then returned to their beaker and FSW was added to approximate a larval concentration of 0.25 larvae mL−1 based on an estimation of the number of remaining larvae. Because larvae of both species of Strongylocentrotus were very small and more individuals were needed for protein estimates, we cultured these species slightly differently. For S. purpuratus, 14 replicate beakers were made so that three samples of 150 larvae each could be taken at 5 dpf from seven of the beakers (the remaining larvae in those beakers were discarded); the other seven beakers were used for 10 dpf samples of 80 larvae each. Thus S. purpuratus is the only species for which the early and mid-development measurements were not paired by beaker. S. fragilis was included in this study opportunistically, so we were not able to repeat the experiment with greater numbers and instead had fewer replicates for S. fragilis than for the other species. S. fragilis had a total of six replicate cultures instead of seven, and the 10 dpf timepoint consisted of only two protein samples rather than three due to a shortage of larvae.
Once larvae were collected, tubes were centrifuged to pellet the larvae and supernatant water was aspirated gently using a pulled pipette. We viewed the expelled aspirated water with a dissection microscope to check for the presence of removed larvae, and on the rare occasions that such larvae were observed the larval count of the sample was adjusted accordingly. If the pellet was disturbed before aspiration was complete, the tube was chilled on ice prior to another round of centrifugation. Once aspiration was complete, the samples were stored at -80 °C for up to three months before protein measurements were made.
To process the protein samples, we removed them from the -80 °C freezer, added 100 µl of 4 °C Nanopure water to each, and placed them on ice. Larval cells were then lysed using Fisherbrand polypropylene pestles designed to fit 1.5 mL microcentrifuge tubes. For consistency in lysing tissue, the pestles were attached to a small handheld screwdriver that spun at 180 rpm; this device was used to grind each sample for 70 seconds. The lysed samples were then returned to the -80 °C freezer for 1-5 days.
A Thermo Scientific Micro BCA Protein Assay Kit was used to determine the protein content of samples against a bovine serum albumin (BSA) standard curve. The samples were removed from the -80 °C freezer, 376 µl of 4 °C Nanopure water was added to each, and they were placed on ice to thaw. Immediately prior to addition to wells, the samples were vortexed for 10-15 seconds. Three subsamples of 150 µl from each homogenized sample were then placed into three experimental wells of a 96-well plate to produce three technical replicates per sample. The number of larvae per technical replicate was then calculated as:
larvae in technical replicate = larvae in original sample * (150 µl technical replicate volume)/(476 µl sample volume)
After addition of reagents and incubation according to the manufacturer’s instructions, the plates were read using a Biotek Synergy H1 Hybrid Microplate Reader. Excel was used to generate the BSA standard curve for conversion of spectrophotometric absorption values to protein content. Two BSA standard ladders were produced per plate. The R2 value of the standard curve exceeded 0.98 each time the assay was performed. The sample readings were then calculated as ng protein per individual for each technical replicate, which were then averaged to yield a single value per sample. The three sample means for each beaker (two, in the case of 10 dpf S. fragilis) were then averaged to yield a single mean ng protein per larva for each beaker.
Ciliated band measurements: To obtain images of larvae for measurement of the ciliated band, 10 larvae were sampled haphazardly from each culture while they were concentrated during the day’s water change. The sampled larvae were placed on a microscope slide, relaxed in 7.5% MgCl2 (made up in tap water) and seawater in a 1:1 ratio, then fixed with a drop of dilute formalin. Larvae were oriented ventral-side up with an eyelash probe, and a coverslip with clay feet was applied. The slide was then viewed with an Olympus BX-51 compound microscope, and the first five correctly oriented larvae encountered were imaged. The microscope was outfitted with a QIClick camera (Teledyne Photometrics) and a motorized z-axis drive, both controlled via MicroManager software. The software enabled us to generate an image stack of each entire larva at 2 µm z-axis intervals.
We later used ImageJ to identify the x, y, and z coordinates of pre-determined landmarks on the ciliated band in images of each larva. A total of twelve landmark maps were used, specific to larvae of each class and developmental timepoint. We calculated the distances between these landmarks to calculate the lengths of short segments of ciliated band, then added these up to yield total ciliated band length (McEdward, 1985; Rendleman et al., 2018).
Analysis: The mean ciliated band length and mean protein content per larva was calculated for each species at each timepoint from the seven (six for S. fragilis) replicate beakers.
The relationship of ciliary band length to protein content was determined for each larval form using a ranged major axis regression, with each species providing two data points, one for each developmental timepoint. Regression lines were estimated using the lmodel2 package version 1.7-3 in R. The 95% confidence intervals slopes and y-intercepts were compared to determine if pluteus and non-pluteus larvae differ in the length of ciliated band they support per unit protein.
* Changed species names to full species names: Astropecten armatus, Apostichopus parvimensis, Dendraster excentricus, Lytechinus pictus, Ophiothrix spiculata, Patiria miniata, Strongylocentrotus purpuratus
* Adjusted parameter names to comply with database requirements
|Age of larva in days post fertilization
|Relative developmental stage of larva
|Beaker that larva was reared in unique for each species
|Identifying number for protein sample. Unique for each Species age and beaker combination
|Average protein content per larva in that sample
|Standard deviation for the protein content estimate for each sample based on three technical replicates
|The northern latitude of the bounding box that includes the collection site of all adult specimens
|The southern latitude of the bounding box that includes the collection site of all adult specimens
|The eastern longitude of the bounding box that includes the collection site of all adult specimens
|The western longitude of the bounding box that includes the collection site of all adult specimens
|The latitude of the laboratory location where experiments took place
|The longitude of the laboratory location where experiments took place
|The year and month (ISO format, yyyy-mm) when the first larval experiment began
|The year and month (ISO format, yyyy-mm) when the final larval experiment concluded
NSF award abstract:
Many ecologically and economically important marine invertebrates (e.g., oysters, crabs, and sea urchins) have life cycles that include feeding larval stages that live drifting in the water as part of the plankton. These larvae spend days or weeks feeding on tiny algal particles to fuel their development until they can metamorphose into juveniles. In nature, however, the plankton includes not only edible particles, but also many particles that are too large to be eaten but which may interfere with feeding on edible particles. These include, for example, large algal particles, eggs and embryos of other invertebrates, re-suspended sediment, and anthropogenic nano- and micro-plastics. When larvae encounter large inedible particles, they may respond by altering their swimming behavior to avoid them, or by capturing and then rejecting them. Such interactions reduce the rate at which larvae can capture edible particles, which forces them to either spend more time feeding before metamorphosis (increasing their overall risk of dying due to planktonic predators), or to metamorphose with less energy, producing juveniles in relatively poor condition. This project examines how large inedible particles affect feeding, time to metamorphosis, and juvenile condition in the larvae of diverse marine invertebrates. The project has the potential to dramatically change our understanding of how larvae feed and survive in natural communities, and thus our understanding of the population dynamics of these important organisms. The project will support research training opportunities for undergraduate and graduate students at California State University Long Beach, a primarily undergraduate institution, as well as summer research internships for students at two local community colleges. Project data will be integrated into laboratory modules in undergraduate courses. Finally, data on the reproductive biology of diverse California marine invertebrates will be added to a public website that is widely used by members of the public, students, and biologists interested in the development, life histories, ecology, and evolution of these common animals.
The factors that control planktonic duration and juvenile condition in marine invertebrates with feeding larvae have long been recognized as critical to understanding their ecology and evolution. Larval feeding environment is clearly one of those factors, but previous work has focused almost exclusively on one feature of that environment, the abundance of food. This project will evaluate the importance of another potentially critical dimension of the larval feeding environment: the presence of large inedible particles, which are frequently abundant in natural plankonic communities. It takes a comparative approach to address two key questions about the effects of large inedible particles on larvae (including those of echinoderms, annelids, and molluscs) that feed using several different particle capture mechanisms. First, do large inedible particles present in natural plankton reduce larval feeding rates? And second, does the presence of large inedible particles extend larval planktonic duration or result in the production of lower quality juveniles? Feeding rates of larvae will be measured in short-term experiments in which larvae are exposed to both food and to natural or artificial large inedible particles over a range of concentrations. Effects of large inedible particles on planktonic duration and juvenile quality will be measured by culturing larvae through their entire life cycles in the presence of large inedible particles at various concentrations. Because feeding performance is an important determinant of planktonic duration, larval survival, and juvenile condition, the project will add greatly to our understanding of how conditions in the plankton affect the population dynamics of the many marine invertebrates with feeding larvae.