Table of Contents

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

Field measurements and bottle collections: Seawater was collected on the research vessel Fay Slover using 5 L Niskin bottles along a gradient from the open ocean near the Chesapeake light tower into the Chesapeake Bay and the James River in June 2019. Conductivity, temperature, and depth were measured using a SBE 32 CTD equipped with a Wetlabs fluorometer and a Wetlabs transmissometer (650 nm).

Field collection of ATP: ATP samples were processed using the hot-water extraction method described by Bochdansky et al. (2021). Three types of ATP samples were collected: 1) pATP in 5 ml seawater filtered through 0.2 mm pore size polycarbonate filters (Isopore GTTP, 25 mm diam.), 2) dissolved ATP (ATP in the filtrate of 0.2 mm filtration), and 3) 0.5 ml ATP in whole water without filtration. Polycarbonate filters were chosen over GF/F filters because of their higher retention efficiency (Taguchi and Laws, 1988; Lee et al., 1995) and because the volumes filtered were small, to ensure filtration times were kept to a minimum even with the use of the smaller pore size. Five milliliters of each depth were immediately and simultaneously filtered using a filtration manifold (25 mm-diameter filters, stainless steel screen supports) preloaded with the polycarbonate filters. The filters were placed in 15 ml polypropylene centrifuge tubes (FalconTM) within seconds after the water passed. The dissolved ATP was captured in 15 ml polypropylene centrifuge vials placed underneath the filtration manifold and inside the vacuum flasks. The dissolved ATP samples (i.e., the 0.2 mm filtrates) were immersed in a boiling water bath for ~15 minutes immediately after filtration to sterilize the water and inactivate ATPases and then frozen at -80 °C until analysis.

In June 2019, instead of boiling on board, filters, filtrates, and whole water samples were collected as above but immediately frozen in liquid nitrogen, transported to the laboratory at -20 °C, and stored at -80 °C in the lab.

Hot-water extraction was then performed on the still frozen samples (i.e., without prior thawing) in the laboratory (see Bochdansky et al. 2021 for details). The shock freezing-boiling treatment breaks up cells more efficiently than boiling alone, which results in higher extraction efficiencies (Bochdansky et al. 2021).

In 2019, 500 ml of water from each bottle was placed in 15 ml centrifuge tubes and shock-frozen in liquid nitrogen. These unfiltered samples thus contained both particulate and dissolved ATP and were labeled total ATP (tATP). All samples were brought to the laboratory in a -20 °C freezer and subsequently stored at -80 °C in the laboratory.

For analysis of the 500 ml shock-frozen samples, boiling hot water was added to the samples and extracted for ~15 minutes in a boiling-water bath.

Particulate and total ATP samples were topped up to 5 ml with ultrapure water using the gradations on the centrifuge tubes and mixed with a vortex mixer. The 500 ml whole-water samples were also diluted to 5 ml with ultrapure water to reduce salt effects that strongly decrease the luminescence yield. It should be noted that the whole-water extraction method used here (for total ATP) requires sufficiently high ATP levels to produce a signal. This was possible because samples were taken in the mesotrophic coastal ocean and in a eutrophic estuary. Such a small amount of water (500 ml) would be insufficient in oligotrophic or deep-sea environments. Hot-water extraction is only one of two methods proposed by Bochdansky et al. (2021). Many of our subsequent collections were based on chemical extraction using phosphorus benzalkonium chloride (P-BAC) instead. Both methods give highly consistent results, with the values from the chemical extraction method exceeding that of the hot water extraction by 20% (Bochdansky et al. 2021). The hot-water method used here has the advantage that measurements of dissolved ATP can be added easily to the protocol as hot water for the inactivation of ATPases is already at hand.

Particulate carbon and nitrogen: During the June 2019 cruise, between 75 ml and 250 ml of seawater (less in the inshore and more in the offshore stations) was filtered onto pre-combusted (450 °C, 4 hours) GF/F filters. The filters were stored frozen (-20 °C) and later dried at 50 °C for approximately 2 days. Filters were then rolled in a tin wrap and pressed into pellets to be analyzed in a Europa 20‐20 isotope ratio mass spectrometer (IRMS) equipped with an automated N and C analyzer.

Laboratory analysis of ATP: Fifty microliters of each sample (in triplicates) were transferred to 6 ml pony scintillation vials (Research Products International), and received 3 ml of ultrapure water, and 50 µL of CellTiter-Glo 2.0 (Promega Corporation). Internal standards were used by spiking a fourth vial with 50 µL of samples with 50 µL of 0.0164 µM ATP standard. Using internal standards instead of separate calibration curves corrects for matrix effects that change the luminescence signal caused by the presence of ions, acids, and organic material (Bochdansky et al. 2021).  Luminescence was analyzed in a PerkinElmer Liquid Scintillation Analyzer with a single photon counting protocol of 1 minute each. The counter was programmed to cycle samples five times in sequence. We determined that values from the second cycle were the most consistent and were thus subsequently used for all analyses.

See supplemental files for formula to calculate ATP, also available in Bochdansky et al. 2021.

Microscopic analysis of biomass, 2019 samples: Prokaryote samples were fixed with formaldehyde and slides were prepared and stained with DAPI as described above. Prokaryotes within an area of 0.00025 mm2 in thirty random fields were manually counted across the filters for each slide under an Olympus epifluorescence microscope. As with the 2018 samples, bacteria counts were converted to carbon using 20 fg per cell.

For eukaryote biomass estimates, 10 ml of the unfixed sample was filtered through a 0.2 µm GTBP membrane filter as well as a 0.45 µm cellulose nitrate backing filter. When no liquid remained, the pump was switched off and the valves were closed to release the vacuum. One ml of a plasma membrane stain (10.0 µL of CellMask (Invitrogen) in 10 ml of 0.2 mm-filtered 34 ppt artificial seawater) was added to the dry filter and kept for 10 minutes before the valves were opened and the pump was turned on. When the stain was completely filtered, two rounds of 1.0 ml filtered artificial seawater were pipetted evenly over the filters to rinse. Nuclei were counter stained with DAPI and embedded in Vectashield as described above. Slides were kept at -20 °C until microscopic examination.

Eukaryote slides were prepared for depths  #1, #3, and #5 at each station (actual depths varied), skipping depths in between (Table 1). Thirty randomly selected image pairs, taken with DAPI and TRITC filter sets, were captured per slide using ToupView Imager software (Fig. 2). DAPI images were taken to confirm that cell nuclei were present. Cell bodies were delineated using ImageJ (Fig. 2) and cell volumes and carbon values calculated as for the Lugol’s samples.

* Split methodology and data processing between 2 datasets. (review from submitter).
* Add sampling date to dataset.
* Adjusted parameter names to comply with database requirements.

NSF Award Abstract:
In the ocean, most living organisms are microbes that are too small to be seen by the naked eye. Despite their small size, microbes play an important role in processes that govern marine ecosystems and food webs. For example, microbes affect the concentrations of nutrients and gases in the water and the atmosphere, thereby exerting a significant impact on the climate globally. Consequently, it is important to know how many microbes there are in any given environment because there is a direct causal connection between living mass and overall biological activity. Determining how “alive” any volume of water is, however, is a difficult task. The gold standard is to count microbial cells under the microscope. This method is extremely time consuming when done well and needs to be performed separately on many different types of microbial cells. In addition, standard microscopic techniques do not reveal whether the cells were alive when they were collected. In contrast, a chemical method based on the amount of adenosine triphosphate (ATP) offers distinct advantages. Notably, ATP is relatively easy to measure, and the method can be widely used because all living cells contain ATP in similar concentrations. This study tests and applies an improved method of ATP analysis to generate data at very high resolution in space and time. One PhD student and six undergraduate students will receive research training and the project fosters international research collaborations with European scientists. This research provides deeper insights into the distribution of live matter in different regions and depths of the world’s oceans.

Decades ago, adenosine triphosphate (ATP) was proposed as a universal biomass indicator. However, its application in the field of oceanography has been limited due to misconceptions regarding cellular ATP concentration. Recent evidence suggests that ATP functions as a hydrotrope requiring homeostatically controlled ATP levels much higher than those solely needed for energy metabolism. ATP occurs in surprisingly stable concentrations in cytoplasm across a wide range of microbes thus representing live cytoplasm volume. This project examines in detail the usefulness of particulate ATP (PATP) as a biomass marker over a large section of the North Atlantic Ocean with special emphasis on mesopelagic and deep-sea environments where chlorophyll is a poor indicator of biomass or associated biological processes. The project uses field collections of marine snow and ambient water in combination with particle cameras to examine the microscale heterogeny of biomass in the water column. Laboratory studies determine factors that may influence the recovery of PATP through filtration and extraction protocols and determine to what extent ATP concentrations potentially deviate from the typical cytoplasm concentration during phosphorus limitation. The improved PATP-biomass method offers numerous operational advantages, especially the fact that it can be employed at high spatial and temporal resolution. Once validated, the PATP biomass method could be widely adopted as a key variable for biomass in routine oceanographic surveys. This project supports graduate and undergraduate students from diverse backgrounds to contribute to laboratory and field research. Public outreach efforts include tours and presentations for middle and high-school students, as well as the general public.

This project is funded by the Chemical Oceanography and Biological Oceanography Programs in the Division of Ocean Sciences.

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.