Facilitation of a tropical seagrass by a chemosymbiotic bivalve increases with environmental stress

Abstract

In fall 2017, we collected sediment, seagrass and lucinids from the east side of Key Biscayne, Florida, USA (25.708822°N, 80.150449°W). Sediment at the collection site is relatively carbonate-rich (53%–60% by acidified weight loss, n = 4) sand with low organic matter content (1.0%–1.7% by loss on ignition, n = 5). Segments of T. testudinum were collected from 1 to 2 m water depth by hand as apical horizontal rhizomes with two to four attached short shoots. Live C. orbicularis were collected from <30 cm sediment depth and separated from sediment in a 4-mm sieve. Sediment was collected adjacent to the seagrass collection area with shovels and sieved over a 2-mm mesh to remove larger material. Organisms and sediment were transported to the Smithsonian Marine Station (SMS) in Fort Pierce and held in aerated tanks with 20 μm-filtered ocean water. C. orbicularis individuals were allowed to burrow in sediment from the collection site for several days until needed for the experiment.

2.2 Experimental design, set up and data collection

We planted T. testudinum segments in 48 custom 25-cm diameter PVC mesocosms consisting of an upper chamber of 14 cm depth and a lower chamber of 3 cm depth, separated by a perforated shelf and porous 100-μm polyester–cotton membrane (Figure S1). The upper chamber was filled with approximately 7 L of homogenized sediment, and the lower chamber was filled with seawater and sealed with a rubber septum. Each mesocosm received a total of nine short shoots, allocated among a 2-shoot, a 3-shoot and a 4-shoot rhizome segment (shoot density 183 m−2). A coloured ziptie tag was placed just in front of the second short shoot on each segment, and the length from the apical end to the tag on each segment was measured prior to planting. Each mesocosm received two 5-cm Rhizon SMS porewater samplers (Rhizosphere Research Products), placed on opposing sides of the mesocosm at approximately 0–5 and 5–10 cm depth.

Mesocosms were housed outdoors in six 416-L aquaculture tanks. Individual tanks were filled with 20 μm-filtered ocean water, aerated with two airlifts and covered loosely with a transparent polycarbonate roofing panel to prevent rainwater input and reduce evaporation. Mesocosm position was re-randomized within tanks during weekly 100% water changes. Epiphytic and microphytobenthic loading were limited. Salinity was maintained at ~34–38 and adjusted downward when necessary with reverse osmosis (RO) water. Surface water pH was ~8.0–8.2, and dissolved oxygen was ~6.3–6.9 mg/L. One submersible aquarium heater (250W; Eheim) was added to each tank at the end of the 5-week acclimation period to stabilize night-time water temperatures.

The experiment was fully factorial, with three factors, eight treatments and six replicates per treatment. There were two levels of each factor: light exposure (ambient-light or shaded), sediment porewater sulphide exposure (ambient or enhanced-sulphide) and C. orbicularis presence (presence or absence of clams). Treatments were arranged in randomized complete blocks, with eight mesocosms in each tank.

Following the 5-week acclimation period, each shaded mesocosm received a single layer of pet-grade window screen held above the canopy by a plastic-coated wire frame. Target average light levels under shading were ~30–100 μmol m−2 s−1, in the range of whole-plant compensation irradiances observed for T. testudinum (Lee, Park, & Kim, 2007). Ambient-light mesocosms received a frame only. Enhanced-sulphide mesocosms were injected with 30 ml of 100 mM sulphide solution, which was prepared with sodium sulphide trihydrate, adjusted to pH 7.1 and injected into the lower chamber of each mesocosm by syringe under anoxic conditions (see also van der Heide et al., 2012). Injections into enhanced-sulphide mesocosms were performed twice at the beginning of the experimental period and weekly thereafter. During the fourth and fifth weeks, the injection regime was adjusted to stabilize the highest sediment porewater sulphide concentrations around 5–6 mM. This target level was chosen in order to reduce the possibility of complete shoot loss in one or more treatments while maintaining a challenge to seagrass sulphide tolerance. Sulphide concentrations of 5–6 mM may induce above-ground die-off in T. testudinum (Ruiz-Halpern, Macko, & Fourqurean, 2008) or may be tolerated if other stressors, such as high temperature, are absent (Koch, Schopmeyer, Kyhn-Hansen, Madden, 2007). Clam-present mesocosms received three randomly selected C. orbicularis, one individual from each available group of 13–17, 19–24 and 25–36 mm shell height. The resulting clam density, 61 m−2, was comparable to the highest field estimates (0–64 m−2, n = 3) at the collection site. Individuals were tagged with bee tags and observed after placement in clam-present mesocosms to ensure burial. Survival throughout the experimental period was 90% (65 of 72 individuals), and all mesocosms retained at least one live C. orbicularis. Neither light nor sulphide exposure significantly affected the number of surviving C. orbicularis in each treatment (two-way Poisson GLM, p > 0.60; Table S1). The condition of live C. orbicularis at the end of the experiment, which was assessed by determining the flesh:shell dry weight ratio (Lucas & Beninger, 1985) after drying for 1 week at 60°C, also did not differ significantly among treatments (two-way ANOVA, p > 0.60; Table S2).

Following acclimation, the experiment was conducted for 6.5 weeks. Temperature and light intensity at canopy level were monitored continuously at 10-min intervals in each tank by two HOBO Pendant loggers (Onset) under ambient-light and shaded mesocosms. Light intensities measured by loggers were converted to estimates of photosynthetically active radiation (PAR) by calibrating loggers to a 4π PAR sensor (LI-193; LI-COR) over 24 hr (Long, Rheuban, Berg, & Zieman, 2012). The average of mean daily PAR over the experimental period across six tanks was 138 μmol m−2 s−1 in ambient-light and 49.6 μmol m−2 s−1 under shades. The average of mean daily temperatures over the experimental period across six tanks was 25.7°C. All calculations excluded data from days involving water changes, major sampling events or other experimental activities with the potential to temporarily but substantially bias recorded light and temperature levels.

Sediment porewater for analysis of total dissolved sulphide (H2S + HS− + S2−) and pH was collected by syringe from both Rhizon samplers in all mesocosms weekly. Porewater was processed the day of collection, prior to sulphide injections in enhanced-sulphide mesocosms. Porewater subsamples (5 ml) were fixed temporarily in an equal volume of sulphide antioxidant buffer (SAOB) to convert all sulphide to S2−, which was measured with an ion-selective silver/sulphide electrode (HI4115; Hanna Instruments). The practical lower reporting limit was considered to be 10 μM. The unfixed remainder of each porewater sample (5–10 ml) was evaluated using a double-junction pH electrode with temperature compensation (S2900C/SAM; Sensorex). Results from the two Rhizon samplers in each mesocosm were averaged to determine mean sulphide concentration and mean pH over 0–10 cm depth.

Samples of sediment porewater for dissolved nutrient analysis (~15 ml) were collected from the 5–10 cm sampler in each mesocosm at initiation and termination of the experiment. Samples of water from each tank were also collected at termination of the experiment, 1 week after the final complete water change. All samples were frozen until analysis. Concentrations of ammonium, ortho-phosphate and nitrate in porewater were determined colorimetrically on autoanalysers (Skalar Analytical; SEAL Analytical) by reaction with salicylate (Kempers & Zweers, 1986), ammonium molybdate (Henriksen, 1965) and sulphanilamide following cadmium reduction to nitrite (Wood, Armstrong, & Richards, 1967) respectively. Water column dissolved nutrient concentrations ranged from 1.8 to 11 μM nitrate, 0 to 6.5 μM ammonium and 0 to 0.004 μM phosphate.

Live shoots in each mesocosm were counted weekly. The percentage net shoot change was calculated as the ratio of the difference of initial and final live shoot counts to the initial live shoot count. One week prior to harvest, all visible shoots were marked with a syringe needle just above the leaf sheath (method modified from Zieman, 1974). New leaf growth was defined as the portion of leaves from the top of the sheath to holes displaced above the sheath.

Rhizome elongation rate was estimated as the difference in length, from planting to harvest, between the apical end and the coloured tag on each rhizome. To account for potential increases in growth with increased above-ground tissue, rhizome elongation rate was standardized to the initial number of short shoots on the segment (i.e. 2, 3 or 4 short shoots) and averaged across the live segments in a mesocosm to estimate a mean rhizome elongation rate for that mesocosm (mm day−1 shoot−1). We also estimated rhizome elongation rates during the acclimation period for several sets of ‘surrogate’ rhizome segments, because our estimates of rhizome elongation in experimental mesocosms included growth that occurred during the acclimation as well as the experimental period. Three groups of three segments (comprising nine short shoots: one 2-shoot, one 3-shoot and one 4-shoot segment) were planted in 7.6-L buckets and maintained in a separate 242-L tank in the same way as the experimental mesocosms during acclimation. Rhizome elongation rates for these surrogate segments were used as an estimate of the range of rhizome growth that may have occurred during the acclimation period in the absence of any experimental treatment. At the end of the experimental period, rhizomes with no attached live shoots were often soft with a strong sulphidic odour.