Refer to the Methods section of Senft-Batoh, et al. (2015).

Collection and Maintenance of Grazers:
Zooplankton from Casco Bay, Maine were collected using a 200 µm mesh net suspended in tidal flow. The catch was transferred to a cooler and transported to the laboratory within 7 hours. Adult female copepods, Eurytemora herdmani, were separated from the sample and placed in a 4 L container filled with 0.2 µm filtered seawater from Long Island Sound, Groton, CT. The copepods (approximately 70 individuals) were maintained at 15°C in a temperature- and light-controlled (12 h:12 h light:dark cycle) chamber and fed a non-limiting diet of the diatom, Thalassiosira weissflogii, and the prasinophyte, Tetraselmis sp. After five days of maintenance in these conditions, adult female copepods to be used in assays were sorted into large petri dishes (10 copepods per dish) containing 80 mL of 0.2 µm filtered seawater. Dishes were set in an environmental chamber (18°C), 24 hours prior to the start of the experiment, to allow the copepods to acclimate to experimental conditions. During this time, the copepods also were starved to ensure complete gut evacuation prior to assays (Dam & Peterson 1988).

Protists were collected from locations in Long Island Sound using a 20 µm mesh net suspended in tidal flow. The heterotrophic dinoflagellate, Polykrikos kofoidii, was isolated from microzooplankton samples collected in Northport Harbor, Long Island, New York. The ciliate, Tiarina fusus, was isolated from samples collected at Avery Point, Groton, Connecticut. The isolates were maintained in 6-well, polycarbonate culture plates. Each well contained 10 mL of a non-limiting mixed diet of the autotrophic dinoflagellates Lingulodinium polyedrum and Scrippsiella trochoidea. This diet is sufficient for the maintenance of both P. kofoidii and T. fusus (Flores et al., 2012). To maintain the microzooplankton populations in culture, subsamples were transferred weekly to new 6-well culture plates containing fresh mixtures of the prey diet. Cultures were kept in a temperature (18°C) and light (12 h:12 h light:dark cycle) controlled incubator. Prior to experiments, Polykrikos kofoidii and Tiarina fusus were micropipetted from the cultures and transferred gently to small petri dishes containing 10 mL of 0.2 µm filtered seawater. Each dish contained 50 individuals. Dishes were returned to the incubator set to 18°C , and microzooplankton were starved for the next 24 hours to ensure complete evacuation of gut contents prior to assays (Tsuda et al., 1989).

Juvenile softshell clams, Mya arenaria (~100 individuals; ~1 cm shell length), were obtained from an aquaculture facility (Downeast Institute for Applied Marine Research and Education) on Beals Island, Maine. Blue mussels, Mytilus edulis (~100 juveniles; ~1 cm shell height), were collected directly from a rocky intertidal outcropping at Avery Point, Groton, Connecticut. Byssal threads of individuals were gently cut to separate them from rocks and from each other. Molluscs were placed in 4 L containers of continuously aerated 0.2 µm filtered seawater in a temperature-controlled (15°C) chamber and fed a non-limiting mixture of Thalassiosira weissflogii and Tetraselmis sp. After 3 days, healthy, actively-feeding clams and mussels were selected for use in experiments. These individuals were moved to 4 L containers of continuously-aerated 0.2 µm filtered seawater and starved for 24 hours prior to assays to ensure complete evacuation of gut contents (Hawkins et al., 1990). Containers were placed in a temperature controlled (18°C) chamber during this 24 hour period to acclimate animals to experimental conditions. Prior to the start of experiments, shell height for mussels and length for clams used in assays was measured. Individuals of near-equal shell height or length were divided among treatments (5 individuals per treatment; mussel mean height= 1.04 (±0.03) cm; clam mean length= 1.07(±0.02) cm). Analysis of variance confirmed that there was no difference in shell height among treatments with mussels (p= 0.232) or shell length among treatments with clams (p= 0.494). Juvenile molluscs were used in the present investigation as juveniles have lower clearance rates than adults; therefore, treatments would not require re-inoculation of the food source as often as would be necessary for adults.

Divers collected ascidians from pilings off of Avery Point, Groton, Connecticut. Individuals of the solitary ascidian, Molgula manhattensis, were removed gently from the substrate. Upon transport to the laboratory, specimens were immediately affixed at the base to a 2 x 2 inch tile of polyvinyl chloride (PVC) with cyanoacrylate glue. The colonial ascidian, Botrylloides violaceus, was found encrusting the shells of live mussels, Mytilus edulis. Colonies cannot be removed from shells without causing significant damage to the organisms. Mussels, therefore, were removed from shells and the valve containing B. violaceus was kept for experiments. Forceps were used to clean the ascidians of any encrusting organisms, such as bryozoans and small arthropods. Ascidians were held in 4 L containers of continuously-aerated, 0.2 µm filtered seawater in a chamber set to 15°C and a 12 h:12 h light:dark cycle. For 2 days, ascidians were fed a non-limiting mixture of Thalassiosira weissflogii and Tetraselmis sp. Individuals then were transferred to 4 L containers of continuously-aerated 0.2 µm filtered seawater and placed in a chamber (18°C) for 24 hours to adjust to experimental conditions.

Direct and indirect induction of toxin production by copepods:
Direct and indirect mechanisms of toxin induction were tested simultaneously using experimental cages. 1 L polycarbonate beakers with bottoms made of 10 µm mesh were nested within another 1 L beaker containing 500 mL of toxic Alexandrium fundyense (300 cells mL⁻¹). The mesh isolated these cells from materials within the cage. Adult female Acartia hudsonica (15 individuals) from either Maine or New Jersey were added to each cage and offered a diet of toxic A. fundyense (300 cells mL⁻¹) or were starved (no addition of algal food). Triplicate treatments (n=3) of the combinations of copepods and algal food within the cages were: 1) Maine copepods fed toxic algae; 2) Maine copepods starved; 3) New Jersey copepods fed toxic algae; 4) New Jersey copepods starved. Control cages (n=3) contained 300 cells mL⁻¹ of toxic algae and no copepods. Assays were run for 72 h, long enough to ensure induction of toxin production, and incubation conditions were identical to those of the algal cultures. Cages were lifted every 12 hours to ensure exchange of cues between compartments. At the termination of the assay, cells of Alexandrium fundyense within cages (where applicable) and below cages were collected from treatments and controls for toxin analysis. Cells within cages (treatments and controls) were enumerated microscopically, before and after incubation, to calculate copepod ingestion rates (Frost 1972). Differences in ingestion rate between the populations were assessed by a t-test.

Culture of Phytoplankton:
The toxic dinoflagellate, Alexandrium fundyense (strain BF-5, isolated from Bay of Fundy, Canada (Anderson et al., 1994)), was grown in semi-continuous culture in F/2 medium without silica (Guillard, 1975). Cultures were maintained in an environmental chamber with fluorescent lighting set to 18°C and a 12 h:12 h light:dark cycle. Toxic cells were used in all assays. Other phytoplankton species used as feed cultures (Thalassiosira weissflogii, Tetraselmis sp., Lingulodinium polyedrum, and Scrippsiella trochoidea) were cultured in the same manner and maintained under the same conditions as Alexandrium fundyense, except that T. weissflogii, a diatom, was grown in F/2 medium containing silica.

Grazer-Induced Toxin Production Assay:
The assay is designed to allow concurrent testing of direct and indirect mechanisms of grazer-induced toxin production on toxin content of Alexandrium fundyense. Separation of direct and indirect mechanisms of stimulation required the use of “cage” vessels. A general description of the experimental design for induction assays can be found in the Appendix. The design allows for manipulation of grazers and algal prey within cages. Such treatments can be used to assess the source of cues by which toxin content is increased. For grazers within cages that are starved (provided with no algal prey), indirect stimulation of toxin production of cue-receiving A. fundyense (300 cells mL⁻¹) below the cages would indicate that a predator kairomone was the cue responsible for increased production. Indirect enhancement of toxin production by grazers actively feeding on toxic A. fundyense suggests that a cue produced by predator interaction with, or digestion of, conspecific cells is a signal for increased toxin production. Increased toxin production in A. fundyense exposed directly to grazers (within cages) could be stimulated by chemical signals but may also involve effects of physical handling of cells or feeding selection by grazers for cells of lower-toxin content.

For each species of grazer, there were three treatments each (n=3) of starved grazers and grazers fed toxic algae. The number of grazers per treatment and the cell density of algae offered as prey (within cages) differed based upon the consumer, and were chosen as to approximate natural abundances and conditions. Control vessels (n=3) containing only toxic A. fundyense, both within and below cages (at the same concentrations as treatments), were prepared to serve as a baseline to detect increased toxin production. Each phylum of grazer was tested in a separate assay with its own preparation of control cells. Indirect induction of toxin production was measured for all species of grazer, but direct induction was only measured for copepod and microzooplankton grazers. The high clearance rates of the shellfish and ascidian species for A. fundyense required re-inoculation of these treatments with algal prey twice daily (12 hours apart). Thus, it was impossible to collect cells that had been directly exposed to grazing from these treatments.

Toxin analysis:
For each assay, extraction of saxitoxins from Alexandrium fundyense proceeded as follows. Cells from treatment and control vessels were collected on a 10 µm mesh and resuspended in filtered seawater in 50 mL centrifuge tubes. Subsamples (1 mL replicates) were drawn from each tube and cells were fixed at a final concentration of 2% Lugol’s solution. Cells were enumerated microscopically to determine the number present in each extract. The 50 mL centrifuge tubes containing A. fundyense were centrifuged at 4,000 x g for 20 minutes. Seawater supernatant was decanted, and the cell pellet was resuspended in 1 mL of 0.1 M acetic acid. Cells were lysed using a probe sonic dismembrator. Sonication was conducted with the tube immersed in ice to prevent heating of the samples. Resulting preparations were centrifuged again at 4,000 x g for 20 minutes to remove cell debris. Next, the acetic acid supernatant (extract) was filtered through a 0.45 µm ultracentrifuge filter cartridge to remove any remaining particulates. Samples were stored at -80°C until analysis. High-performance liquid chromatography with fluorescence detection (HPLC-FD; Oshima, 1995) was used to identify the carbamoyl paralytic shellfish toxin derivatives of saxitoxin (STX), neosaxitoxin (NEO), and gonyautoxins 1 through 4 (GTX 1-4), as well as the less toxic sulfamate congeners, C1 and C2 (Indrasena and Gill, 1998). Toxins were quantified by comparison to known standards (Certified Reference Materials Program, NRC Institute for Marine Biosciences, Canada) and expressed as mass of STX equivalents per cell according to conversion factors of Oshima (1995).

senft-batoh_2015b.csv