Table of Contents

• Coverage
• Dataset Description
  • Methods & Sampling
  • Data Processing Description
  • BCO-DMO Processing Description
• Data Files
• Related Publications
• Parameters
• Instruments
• Project Information
• Funding

Collection Media and inoculum:

Sponge exhalent water and reef sea water, for use as media in the incubation experiment, were collected on 9-December-2020 from Looe Key Reef Sanctuary Preservation Area (24.54605, -81.40610) in the Florida Keys National Marine Sanctuary (FKNMS). Sponge exhalent water was collected from each of 4 individuals of two sponge species, Niphates digitatalis and Xestospongia muta. Exhalent water collection was completed by scuba divers using a three-way valve with one valve attached to a ~8cm piece of PharMed L/S 25 tubing (Masterflex® L/S® Precision Pump Tubing, Radnor, PA, USA) that was placed into the excurrent osculum of the sponge to collect exhalent water, a second valve attached to a syringe to create suction and pull in exhalent water, and the final valve attached to ~8cm piece of Teflon tubing that further attaches to the stainless-steel fitting on a 1 L FlexFoil PLUS bag (SKC, Eighty Four, PA, USA) for final exhalent water storage. Exhalent water was pulled into the syringe at a rate of ~2ml min-1, below the estimated pumping rate of both sponge species. Two bags (~1L) were filled per sponge and all sponges were located within 4 meters of one another at ~8m depth. Upon return to the boat all bags were sealed and placed in an ice-filled cooler until further processing in the lab (~2 hrs). 

Reef surface water, for use as both reef metabolome media and picoplankton inoculum during the incubation experiment, was collected in 6 acid-washed 2 L Nalgene© HDPE bottles (Thermo Fisher Scientific, Waltham, MA, USA) from ~1 m below the surface just above the reef at Looe Key. Metabolome bottles were immediately placed in a cooler filled with ice until return to the lab (~1 hr). The reef picoplankton inoculum was kept in the dark at ambient seawater temperature.

Picoplankton incubation experimental setup:

The 48-hour experiment included two treatments, reef (i.e., control) and sponge metabolome, that were sampled at 0 hours, 24 hours, and 48 hours (T0, T24, T48, respectively). Surface seawater (6 L) for picoplankton inoculum was filtered through three 1.6 µm, 47 mm pre-combusted GF/A filter to remove larger eukaryotic plankton and particular matter. Filters were housed in a 47 mm acid-washed in-line PFA filter holder (Advantec, Cole-Parmer, Vernon Hills, IL, USA). Filtering was completed via peristalsis (MasterFlex L/S pump and pump heads, Cole-Parmer, Vernon Hills, IL, USA). Samples were filtered slowly (50 ml min-1) to avoid bursting of microbial cells on the filter membrane and filters were changed every 2 L. PharMed L/S 25 (Masterflex® L/S® Precision Pump Tubing, Radnor, PA, USA) acid-washed tubing was used for all filtration. The sponge and reef metabolome media (6 L each) were both passed through 0.22 µm, 47 mm polytetrafluoroethylene (PTFE) filters (Omnipore, EMD Millipore Corporation, Billerica, MA, USA) using the same process detailed above. Sponge metabolome media consisted of a 3:1 ratio of filtered sponge exhalent water (1:1 exhalent water from X. muta and N. digitalis) to 0.22 µm filtered sea water.

Following filtering, two 20 L food grade containers were used, one per treatment, to pool the filtered inoculum with the filtered media. Pooling was done to ensure consistent starting mixtures for each incubation bottle. Following initial (T0) sampling as detailed below, pooled treatments were evenly distributed into four 2 L Nalgene© bottles per treatment for an approximate ratio of 1:2 filtered picoplankton inoculum to filtered media. All incubation bottles were kept in a dark incubator at 26ºC for the duration of the 48-hr experiment and only removed for picoplankton community sampling at T24. Water chemistry and cell abundance was assessed for both the sponge and reef metabolome treatments at the start (T0) and end (T48) of the incubation experiment. Samples were also taken at T24, but they were only used to obtain cell abundance.

Flow cytometry

To quantify picoplankton abundances, 1 ml samples of water from each pooled treatment (T0; n=3 per treatment) and from each incubation bottle (n=4 per treatment) at both T24 and T48 was collected in a sterile cryovial and fixed with paraformaldehyde (1% final volume), incubated in the dark at 4ºC for 1-hr, and then frozen at -80ºC. The fixed samples were used for enumeration of total bacterial cells (Prochlorococcus, Synechococcus, picoeukaryotic cells, and unpigmented cells (heterotrophic bacteria and archaea)) via flow cytometry at Bigelow Laboratory for Ocean Sciences. Flow cytometry analysis was completed using a Bio-Rad ZE5 (Hercules, California, USA) with a 488 nm laser activated. Samples were thawed and pre-screened through 70 µm mesh and diluted 1:10 with Tris EDTA (TE) Buffer pH 8.0. Cells were then stained using 10x working stock DNA stain SYBRGreen I (Thermo Fisher Scientific, Waltham, MA, USA) following the protocol of Marie et al. (2005). A total of 180 µl of diluted sample was run at a flow rate of 0.5 µl sec -1. Particles were excited with the 488 nm blue later and data acquisition was triggered by green fluorescence. Data signals were recorded by detectors with three bandpass filters including forward scatter (FSC), right angle light scatter (SSC), and fluorescence emission in green (525/35nm). To minimize noise, samples were subgated in FSC-405. Data files logarithmic scatter plots of fluorescent and light scattering properties were analyzed and total bacterial counts were identified based on size and presence of the green fluorescence. All counts were converted to cell abundance based on sample volume (including adjustments for preservation, dilution, and staining).

Biochemical Analyses

Total Organic Carbon (TOC) and Total Nitrogen (TN)

For TOC and TN analyses, 20 ml of water was collected from the pooled treatments prior to their distribution into individual incubation bottles (T0: reef metabolome, n=3; sponge metabolome, n=3) and for each bottle at the end of the experiment (T48: n=4 per treatment) in acid-washed, combusted 40-ml amber EPA vials. Samples were immediately acidified to pH 3 using 12M trace-metal grade hydrochloroic acid (HCL, OptimaTM, Fisher Chemical, Fisher Scientific, Hampton, NH, USA) and stored in the dark at 4ºC. The acidified samples were processed at Woods Hole Oceanographic Institute using a Shimadzu TOC-L analyzer (Longnecker et al. 2015). Additionally,

Filtering for Biochemical Analyses

All other biochemical analyses were completed only on filtered water samples from the pooled treatments at T0 (reef metabolome, n=1 and sponge metabolome, n=2) and T48 incubation bottles (n=4 per treatment). Approximately 680 ml of water from each incubation bottle was individually passed through a 0.22 µm, 47 mm PTFE filter using peristalsis as previous detailed except for a slightly slower filtering rate (30 ml/min) to avoid bursting the microbial cells. Following filtration, the PTFE filters were placed in individual sterile cryovials and stored at -80ºC. 

Inorganic macronutrients

For inorganic macronutrient analyses (PO43-, NO3-, Silicate, NO2, NH4+), ~25 ml of the 0.22 µm filtrate from each sample was collected in an acid-washed 30 ml break-proof Nalgene© HDPE bottle (Thermo Fisher Scientific, Waltham, MA, USA), frozen and stored at -20ºC before being shipped to College of Earth, Ocean, and Atmospheric Sciences at Oregon State University for inorganic macronutrient analyses. Phosphate (PO43-) and ammonium (NH4+) were measured using a Technicon Auto Analyzer II (SEAL Analytical Inc., Mequon, Wisconsin, USA) and silicic acid (i.e., silicate), nitrate (NO3-) and nitrate+nitrite (NO3- + NO2-) were measured using an Alpkem RFA 300 colorimetric autoanalyzer (Alpkem, Kranj, Slovenia). Analytical methods and data processing were completed as described in Gordon et al. (1994), but a brief summary of each analysis is provided. The PO43- method was modified from the molybdenum blue procedure (Bernhardt and Wilhelms 1967), in which PO43- is determined as reduced phosphomolybdic acid employing hydrazine as the reductant. The indophenol blue method, modified from ALPKEM RFA methodology, was used for the measurement of NH4+. Silicic acid was measured using the methodology of Atlas et al. (1971) in which the addition of an acidic molybdate reagent forms silicomolybdic acid that is then reduced by stannous chloride. Lastly, NO3- and NO3-+ NO2- were measured based on the methods of Atlas et al. (1967) with modifications to improve precision. Sulfanilamide and N-(1-Napthyl)ethylenediamine dihydrochloride react with NO2- to form a diazo compound. For the NO3- + NO2- analysis, NO3- is first reduced to NO2- using an OTCR and imidazole buffer as described by Patton (1983). The NO2- analysis is performed on a separate channel, omitting the cadmium reductor and the buffer.

Dissolved combined neutral sugars (DCNS)

Dissolved combined neutral sugars (DCNS) were measured from frozen 20 µl aliquots of 0.22 µm filtered seawater (collected in acid-washed, combusted 40-ml amber EPA vials and stored in the dark at 4ºC) at the Complex Carbohydrate Research Center, University of Georgia. Each sample was first desalted and hydrolyzed. For desalting, each sample was loaded onto a gravity column, containing forty milliliters of mixed ion exchange resins (AG 501-X8, 20-50 mesh, Bio-Rad), that was packed and prewashed with 200 ml (5x bed volume) of nano-pure water. Samples were then eluted with 120 ml (3x bed volume) of nano-pure water. The resulting flow-through and wash solution were lyophilized. Following lyopholization, the recovered materials were hydrolyzed wth 2 ml of 2 N TFA at 100ºC until high-performance anion exchange chromatography (HPAEC) analysis. By employing a specific HPAEC program, as detailed below, the neutral monosaccharides can be separated allowing the measurement of carbohydrates in each sample.

Monosaccharide standards, including fucose (Fuc), rhamnose (Rha), arabinose (Ara), glucose (Glc), galactose (Gal), xylose (Xyl), mannose (Man), and fructose (Frc), were hydrolyzed in the same manner and at the same time as the samples. Three concentrations of the standard mixture were prepared serially to establish a calibration equation. The quantity of each residue in the sample was calculated by linear interpolation of respective residue area units into the calibration equation.

Monosaccharides from each sample were analyzed by HPAEC with pulsed amperometric detection (HPAEC-PAD) using a DIONEX ICS3000 system (Thermo Fisher Scientific) equipped with a gradient pump and an electrochemical detector. The carbohydrates were separated by a Dionex CarboPac PA20 (3x150mm) analytical column with an amino trap column and eluted with degassed 12 mM NaOH. Injections were made every 40 min. Under the HPAEC conditons, Xyl and Man cannot be separated (Borch and Kirchman 1997) and are presented as combined results. Samples were analyzed in triplicate and mean values were reported as ng ml-1 based on the volume analyzed.

Fluorescent dissolved organic matter (fDOM)

Flourescent dissolved organic matter (fDOM) was measured from cool (4ºC) aliquots of 0.22 µm filtered seawater (collected in acid-washed, combusted 40-ml amber EPA vials and stored in the dark at 4ºC) from each sample following the methods detailed in Nelson et al. (2015). Samples were analyzed using a Horiba Aqualog scanning fluorometer with 150 W Xe excitation lamp, Peltier-cooled CCD emission detector, and a simultaneous absorbance spectrometer (Horiba Scientific, Piscataway, New Jersey, USA). Samples were brought to room temperature and loaded into DIW-leached and rinsed quartz cuvettes (1 cm diameter). Excitation-emission matrices (EEMs), 3D contour plots of excitation and emission fluorescence, were measured for each sample. Analysis began and ended with 4 DIW-filled cuvettes as blanks. EEMs were processed with a MATLAB script (https://github.com/zquinlan/fDOMmatlab/script.m) that employs parallel factor analysis (PARAFAC) to identify peaks that correspond to previously characterized fDOM components (Coble 1996, Nelson et al. 2015).

Targeted metabolomics

For targeted metabolomics analysis, the remaining filtrate from each sample (~600 ml) was acidified to pH ~3 using 12M trace-metal grade hydrochloric acid (HCL) and the volume was recorded. Note that for targeted metabolomics analysis only one T0 sample from each treatment was analyzed. Extracellular metabolites were concentrated and extracted from each acidified filtrate using solid phase extraction (SPE) with 1g/6cc BondElut PPL cartridges (Agilent, Santa Clara, CA, USA) using a vacuum manifold (Waters Corporation, Milford, MA, USA) following the protocols from Kido Soul et al. (2015) and Fiore et al. (2017). Briefly, the acidified filtrate from each sample is passed through acid-washed FEP tubing and a pre-conditioned (with 100% HPLC-grade methanol) 1g/6cc BondElut PPL cartridges (Agilent, Santa Clara, CA, USA). Following filtration, each cartridge is wrapped in combusted aluminum foil, placed in a labeled, sterile Whirl-Pak bag and frozen at -80ºC. Frozen cartridges were shipped to Appalachian State University where the extraction process was completed by rinsing each thawed cartridge with 4 volumes of 0.1 M HCL followed by a gentle 5 min drying cycle using vacuum pressure and finally eluted into acid-washed, combusted 8-ml glass vials with 6 ml of 100% methanol. Using combusted glass pipettes, the methanol extracts were transferred to acid-washed, combusted 8-ml amber EMP vials and dried down to a single droplet using vacuum centrifuged then stored at -20ºC. 

Targeted metabolite analysis was performed at Woods Hole Oceanographic Institute using UPLC (Accela Open Autosampler and Accela 1250 Pump, Thermo Scientific) coupled with a heated electrospray ionization source (H-ESI) and a triple stage quadrupole mass spectrometer (TSQ Vantage, Thermo Fisher Scientific) in selective reaction monitoring (SRM) mode per methods detailed in Kido Soule et al. (2015). The raw XCalibur files were converted to mzML files using msConvert (Chambers et al. 2012) and processed with MAVEN (Melamud et al. 2010) to obtain calibration curves based on the integrated peak area generated for each metabolite. To calculate environmental concentrations the concentration of the metabolite was divided by the volume of the original acidified filtrate passed through the PPL cartridge. Lastly, metabolites that met the threshold detection and quantification limits for the targeted analysis were corrected for extraction efficiency by dividing their environmental concentrations by their published extraction efficiency in sea water (Johnson et al. 2017).

* Adjusted parameter names to comply with database requirements.

NSF Award Abstract:
The seawater around coral reefs is typically low in nutrients, yet coral reefs are teeming with life and are often compared to oases in a desert. Life exists in these 'marine deserts' in large part, due to symbiotic associations between single-celled microbes and invertebrates such as corals and sponges. The concentration and type of dissolved organic matter (DOM), a complex pool of organic nutrients such as amino acids, vitamins, and other diverse compounds, also affects the health of coral reefs. The composition of DOM on coral reefs is linked to both the composition of free-living microbes in the seawater and to the nutrition of filter-feeding organisms, such as corals and sponges. However, the factors that influence the composition of DOM on coral reefs and the consequences of how it changes are not well understood. Recent work suggests that sponges could have a significant impact on the composition of reef dissolved organic nutrients, depending on sponge species due to differences in filtration capacity and in their symbiotic microbial communities. This project characterizes how diverse sponge species process DOM on coral reefs and determines the impacts of this processing on the free-living microbial community. Seawater is collected from sponges (pre- and post- sponge filtration) on coral reefs in the relatively pristine region of Curacao, and incubation experiments measure the impact of sponge filtration on the growth of the free-living microbial community. The organic nutrients of seawater samples are analyzed using cutting-edge techniques to distinguish the types of nutrients that are processed by sponges. The incubation experiments, using free-living microbes collected from the coral reef, quantify the impact of sponge filtration on the growth and composition of this community. This project provides fundamental understanding of how sponges contribute to the base of the coral reef food web. As the human-driven impacts continue to alter the composition of organisms on reefs, this understanding is necessary to predict changes to reef microbial food webs and is thus essential for scientists, reef managers, and policy decision makers. This project trains undergraduate students and a postdoctoral scholar and contributes to undergraduate and K-12 education through development of sponge-centric lessons that focus on local U.S. east coast aquatic environments as well as coral reef ecosystems.

Sponges vary in their capacity to filter seawater and in their associated microbial communities, leading to diverse metabolic strategies that often coexist in one habitat. While it is well-established that sponges are important in processing dissolved organic matter (DOM), an important reservoir of reduced carbon compounds, and transferring this energy to benthic food webs, there has been limited work to understand the consequences of sponge processing on the composition of coral reef DOM and on pelagic food webs. Specifically, while studies have shown that exudates of corals and algae select for specific groups of picoplankton (autotrophic and heterotrophic, respectively), similar data for sponges are required to understand the multiple factors that shape the composition of DOM and of the picoplankton community on coral reefs. Thus, this project is aimed at addressing a major knowledge gap of the role of sponge-derived DOM (sponge exometabolome) in coral reef biogeochemistry. An in situ sampling design targeting prominent Caribbean sponges and picoplankton incubation experiments is coupled to address both the composition of sponge exometabolomes and delineate shifts in the picoplankton community derived from sponge exometabolomes. Molecular-level changes to seawater DOM by sponge processing and the impact of these changes on the overall coral reef DOM profile is assessed with two DOM analysis techniques: a commonly used fluorometry technique (fDOM analysis) and with high-resolution mass spectrometry (LC-MS/MS). Additionally, microbiome and functional gene profiling, growth metrics, and nutrient analyses are employed to assess changes in the picoplankton community in response to sponge exometabolomes. Advanced data analysis techniques then synthesize data generated by each approach to provide novel insight on a poorly uncharacterized biogeochemical pathway on coral reefs. The work outlined here represents entirely novel information on the impact of sponge metabolism on the composition of DOM, sheds light on biologically important molecules involved in benthic-pelagic coupling, and importantly, generates data using standardized methods, thus facilitating comparison to previous and future DOM datasets.

This award reflects NSF's statutory mission and has been deemed worthy of support through evaluation using the Foundation's intellectual merit and broader impacts review criteria.