Dataset: Lateral line development in Atlantic silverside (Menidia menidia) larvae reared at contrasting pCO2 conditions in a laboratory experiment conducted with embryos from adults collected in Long Island Sound in 2021

Spawning Atlantic silversides (Menidia menidia) were caught on May 24th, 2021 in Mumford Cove (41°19’25”N 72°01’07”W). Fish were captured using a beach seine (30 × 2m) and were transported to the University of Connecticut’s Rankin Seawater Laboratory, separated by sex, and held at 20°C in 80-l aerated tanks. The following day, fish were strip-spawned (following Murray et al., 2014, 2016; Malvezzi et al., 2015), and within one hour embryos were randomly distributed into 20-l replicate rearing containers. 

Experimental Design:

Each container was equipped with 4 mesh-covered holes (7 cm diameter, 150 µm mesh) to allow water exchange with the individual surrounding 600-l tanks in the Automatic Larval Fish Rearing System (ALFiRiS). This custom-built system consists of nine such recirculation units that are independently controlled for pH and temperature via a customized LabView program (Baumann et al., 2022; Zavell and Baumann, 2024). ALFiRiS measures conditions once per hour via sampling pumps connected to a central pH sensor (Hach pHD digital electrode) that is calibrated weekly using NIST 2-point pH references. As needed, the system automatically adjusts pH /pCO_² levels by triggering solenoid valves to inject standardized amounts of 100% bone dry CO₂ (to decrease pH) or CO₂-stripped air (to increase pH), while maintaining desired temperatures via computer-controlled submersible heaters (Finnex Deluxe Titanium, 800W) or in-line chillers (DeltaStar ⅓ horsepower). All trials experiments were conducted at 22°C to resemble late spring conditions at Mumford Cove normally experienced by spawning silversides and their larvae (Murray and Baumann, 2018).

A  lateral line development experiment  specifically tested our hypothesis that high CO₂ environments affect schooling due to morphological changes and/or the development of the LL system. In both experiments three CO₂ treatments were used, referred to hereafter as control (8.15 pH units, ~400 µatm CO₂), elevated (7.45pH units, ~2200 µatm CO₂), or extreme (7.20 pH units, ~4200 µatm CO₂) CO₂ conditions. The control CO₂ treatment is characteristic of the average local surface ocean and of coastal systems at the start of the silverside spawning season in spring (Murray et al., 2014). The elevated CO₂ level represents the maximum predicted open ocean CO₂ for the next 300 years (Caldeira and Wickett, 2005), and it is both a common benchmark in ocean acidification studies as well as a level already observed in productive coastal systems during summer (Baumann et al., 2015). The extreme CO₂ level is rarely reached yet in contemporary systems (Baumann et al., 2015), but may become more common in productive nearshore habitats under future climate and eutrophication scenarios (Wallace et al., 2014). 

Actual CO₂ levels were determined biweekly by measuring total alkalinity (A_T; Murray and Baumann, 2018) in 300 mL samples of filtered seawater (10 µm) taken from each tank. Salinity was measured concurrently using a handheld refractometer. Endpoint titrations (G20 Potentiometric Titrator, Mettler Toledo) were calibrated with Dr. Andrew Dickson’s certified reference material for AT in seawater (UCSD Scripps Institution of Marine Science; Dickson et al., 2003) and titration accuracy was (±1%). Measured temperature, salinity, pH, and AT were then used to calculate CO2 (CO2SYS V2.1; https://cdiac.ess-dive.lbl.gov/ftp/co2sys/CO2SYS_calc_XLS_v2.1/) using K1, K2 constants from Mehrbach et al. (1973), refitted by Dickson & Millero (1987), and Dickson (1990) for potassium bisulfate (KHSO4). 

Upon hatch, larvae were fed both a powdered weaning diet (Otohime Marine Fish Diet, size A1, Reed Mariculture, Campbell, CA, USA) and newly hatched brine shrimp nauplii ad libitum (Artemia salinia, San Francisco Strain, brineshrimpdirect.com, Ogdon, UT, USA). During the remainder of each experiment larvae were fed daily ad libitum rations of newly hatched nauplii. Ammonia was measured daily (API Saltwater Test Kit) to assure that levels remained < 0.25 ppm. Rearing containers were siphoned daily to remove uneaten food and detritus, and 25% partial water changes were conducted weekly. Salinity (27 – 31 psu) and light conditions (15 h light: 9 h dark) reflected optimal levels for growth for this species (Middaugh et al., 1987). Throughout both experiments, sampled larvae were euthanized and fixed in cold (2-4°C) 4% paraformaldehyde (PFA) in phosphate buffered saline (PBS). 48 hours following fixation, larvae were transferred to 70% ethanol in deionized water, which was changed twice to remove excess PFA.

Lateral Line Development Experiment:

The lateral line development experiment quantified the rate of development of the cranial lateral line system including the number of neuromasts under contrasting CO₂ conditions (control and extreme, as defined above). The cranial lateral line system (as opposed to the trunk LL system) was studied given its role in maintaining spatial positioning within a school of fish (Mekdara et al., 2021). Approximately 500 embryos were randomly allocated to each of two 20-l rearing containers (one control and one extreme CO2 treatment). Hatching occurred 9 days post fertilization (dpf; June 1st, 2021), and 15 larvae were sampled daily from 0 to 7 dph, and then 3× per week until 34 dph (to ~14 mm Standard length of the sampled larva (SL)). Each day 5 larvae were used for vital fluorescent staining of the neuromasts using 4-Di-2-ASP 4-(4-Diethylaminostyryl)-1-methylpyridinium iodide(4-Di-2-ASP; Nakae et al., 2012; Nickles et al., 2020) and the other 10 sampled larvae were euthanized and fixed as previously described for future imaging of neuromasts using SEM (scanning electron microscopy).   

Fluorescent imaging of stained larvae was done on a Nikon SMZ 1500 dissecting microscope with  epifluorescence (GFP filter set) and captured with a SPOT® digital camera and SPOT software (v 5.2.21, Diagnostic Instruments Inc., Sterling Heights, MI). Then larvae were fixed as previously described and subsequently measured (notochordal length, NL, for pre-flexion larvae; or  SL, for post-flexion larvae; and total length, TL). Adobe Photoshop® CS3 was used to generate panoramas from multiple images to visualize complete distribution of neuromasts. Adobe Illustrator® CS3 was then used to make diagrams of neuromast distributions from panoramas. Neuromast numbers were determined for each larva by two independent analysts blinded to the treatments. 

Scanning electron microscopy (SEM) was used to determine the degree of canal formation, in larvae in the control (ambient) and extreme groups between 0 and 34 dph (nAmbient = 22; nExtreme = 22). Larvae ranged in size from 5 to 16 mm NL with a mean (± SE) of 9.3 ± 0.7 and 10.3 ± 0.5 mm in the control and extreme CO2 treatments, respectively. Larvae were dehydrated in an ascending series of ethanol solutions at 1–2-hour intervals (50%, 75% [overnight], 85%, 95%, 100%), critical-point-dried in liquid CO₂ (Leica CPD 300), mounted on aluminum stubs with adhesive carbon discs, and sputter-coated with platinum (Leica MED 020, 15 nm). Imaging was done at 3 kV at a working distance of 10-16 mm on a Zeiss Supra 40VP SEM at the Marine Biological Laboratory (Woods Hole, MA). The degree of canal formation in the vicinity of each neuromast (a canal segment) was assigned to one of four developmental stages (Tarby and Webb, 2003; Webb and Shirey, 2003). Stage 1 - superficial neuromast on epidermis, Stage 2 - neuromast sunken into a groove of unossified or ossified canal walls (could not be distinguished in SEM’s), Stage 3 - epithelium fused over the neuromast forming a canal roof, and Stage 4 - bony canal wall fused over the neuromast. Complete ossification of the bony canal (stage 4) could only be quantified using histology, so canal segments that were enclosed with a canal roof were classified as stage 3/4. 

Organism Identifier (LSID, Life Sciences Identifier):

Menidia menidia, urn:lsid:marinespecies.org:taxname:159228