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Major project objectives are as follows - (1) use isotope and other tracer methods to examine the major sources of nutrients, carbon, and sulfur to the south Florida ecosystem, (2) use geochemical methods to examine the major forms of nutrients, carbon, and sulfur in the sediments, the stabilities of the observed chemical species, and sinks of these elements in the sediments, (3) examine the biogeochemical processes controlling the cycling of nutrients, carbon, and sulfur in the ecosystem, and use geochemical modeling of porewater and sediment chemical data to determine the rates of these recycling processes, (4) develop geochemical sediment budgets for nutrients, carbon, and sulfur on a regional scale, including accumulation rates of these elements in the sediments, fluxes out of the sediments, and sequestration rates, (5) collaborate with mercury projects (USGS ACME team and others) to examine the role of sulfur and sulfate reduction in the production of methyl mercury in wetlands of south Florida, and the bioaccumulation of mercury in fish and other wildlife, (6) develop a geochemical history of the south Florida ecosystem from an examination of changes downcore in the concentration, speciation, and isotopic composition of nutrients, carbon and sulfur; use organic marker compounds and stable isotopes to develop a model of seagrass history in Florida Bay, (7) incorporate information from nutrient studies in overall ecosystem nutrient model, and results from sulfur studies in ecosystem mercury model.
Studies of sulfur within the ecosystem relate directly to the issue of methyl mercury production and bioaccumulation, a serious threat to both wildlife and to the human population. Microbial sulfate reduction in wetlands (an anaerobic process) is the primary driver of mercury methylation. Understanding the source of sulfate to the wetlands of south Florida may be a key to understanding why mercury methylation rates are so high, and on how remediation efforts in the Everglades may impact mercury methylation rates. We are also examining the sulfur geochemistry of sediments on a regional scale, with emphasis on areas that are methyl mercury "hotspots". We are emphasizing co-sampling with USGS mercury researchers (ACME team).
The geochemical history component of this project will provide information on historical changes in the chemical conditions existing in south Florida wetlands. This will provide wetland managers with baseline information on the water quality goals needed to achieve "restoration" of the ecosystem. It will also provide land managers with an estimate of the range of water quality and environmental conditions that have affected the south Florida ecosystem in the past. Geochemical history data in combination with information from paleontologic studies of the USGS paleoecology group and others will also provide insights on how organisms in the south Florida ecosystem have responded to environmental change in the past, and predict how these organisms will likely respond to changes in the ecosystem resulting from restoration efforts.
Kotra, R. K.; Holmes, C. W.; Orem, W. H.; Hageman, P. L.; Briggs, P. H.; Meier, A. L.; Brown, Z. A.
Spiker, Elliott C.; Holmes, Charles W.
Holmes, C. W.; Kendall. C.; Lerch, H. E.; Bates, A. L.; Silva, S. R.; Boylan, A.; Corum, M.; Marot, M.; Hedgman, C.
Lerch, H. E.; Rawlik, P.
Total carbon, total nitrogen, and total sulfur contents of the lyophilized sediments and seagrass fragments were determined using a Leco 932 CHNS Analyzer. Organic carbon contents were determined on the Leco 932 CHNS Analyzer after treatment of the samples in acid to remove carbonates. We used an acid fuming method adapted from Hedges and Stern (1984) and Yamainuro and Kayanne (1995) to remove carbonates. Our procedure involved: (1) weighing of the sample into silver Leco sampling cups on a microbalance, (2) placing the weighed silver cups in a sealed chamber (dessicator) with concentrated HCl in the bottom, (3) allowing a minimum of 72 hours for the acid fuming to remove all carbonates from these carbonate-rich sediments and seagrass fragments, and (4) redrying and reweighing the cups prior to analysis.
Total phosphorus was determined by a method adapted from that of Aspila et al. (1976). Lyophilized sediment was weighed into crucibles and baked for 2 hrs. at 55°C. The baked sediment was cooled and quantitatively dumped into 250 ml plastic centrifuge cones containing 50 ml of 1M HCl. The baked sediment was extracted on a shaker in the 1M HCl for 16 hrs. An aliquot of each extract was centrifuge filtered through 0.45 micron centrifuge filters, and the filtrate adjusted to pH 7 with NaOH and transferred to plastic test tubes. The filtrate was then analyzed for phosphate using the standard phospho-molybdate calorimetric method (Strickland and Parsons 1972).
The stable isotopic composition (delta C and delta N) of selected sediments and seagrass fragments was determined using a Micromass Optima continuous flow mass spectrometer coupled to a Carlo Erba elemental analyzer. Delta 15N was determined on whole sediment or seagrass samples, while delta 13C was determined on sediment and seagrass fragments after acid fuming of the samples, as described above.
Analytical precision (percentage relative standard deviation) for the elemental analysis of sediments and seagrass fragments varied from sample to sample, but generally was as follows: 2% for total carbon, 1% for total nitrogen, 10% for total sulfur, 3% for total phosphorus, and 4% for organic carbon. Stable isotope analysis of the fine sediment had an analytical precicion (1 sigma) of about 0.1 to 0.2 per mil for both delta 13C and delta 15N, but the precision for seagrass fragments was as high as 0.5 per mil due to sample heterogeneity.
Porewater Analysis Sediment porewater was obtained from cores using an in situ squeezing technique described in detail elsewhere (Orem et al. 1997). In brief a piston core is taken using an acrylic core tubewith a series of threaded ports at intervals along its length. The ports are closed during coring with screws and small O-rings. After the core is collected, it is returned to a dry land site (parking lot at hotel, etc.) and bolted onto a squeezer board. The squeezer board consists of a vertical metal frame with threaded metal plates attached to the frame, threaded rod through the plates, and pusher pistons at the ends of the rods. The core barrel containing the sample already has a piston at the top used for coring, and a second piston is inserted at the bottom. The threaded rods with pusher pistons are advanced through the threaded plates with a ratchet until they make contact with the pistons in the core barrel at both the top and bottom. Depth intervals in the core are selected, and the screws and O-rings in the threaded ports are removed and replaced with a threaded fitting. The fitting is threaded into the port and has a piece of tubing on the inside which extends to the middle of the core. Thus, during squeezing only porewater from the center of the core is collected. The fitting has a luer lock on the outside on which a syringe filter (0.45 micron) is attached. The syringe filter is attached to a collection bottle or a syringe by a shorth piece of tubing. Typically, 10-14 ports are selected downcore for porewater sampling, with close interval sampling near the surface and greater spacing of intervals at depth. Squeezing is initiated by turning the threaded rods with a ratchet so that the core is compressed by the pistons. After squeezing is initiated, most squeezing is done from the bottom to preserve the integrity of the core near the surface. Porewater is forced into the tube at the center of the core and exits the core through the syringe filter and is collected. Porewater yields obtained range from none in some dry holes to more than 100 ml. Florida Bay sediments are more difficult to squeeze than peat from the Everglades due to the fine-grained nature of the carbonate muds and their incompressibility compared to peat. Typical yields from Florida Bay sediments was 10-20 ml of porewater.
Porewater was analyzed for the following parameters where sufficient volume was available: phosphate, ammonium, chloride, fluoride, sulfate, sulfide, redox, pH, titration alkalinity, conductivity, salinity, and metals. Phosphate and ammonium were analyzed calorimetrically after removal of sulfides (Strickland and Parsons 1972). Chloride, fluoride, and sulfate were analyzed by ion chromatography. Sulfide, redox, conductivity, salinity, pH, and titration alkalinity were determined by electrochemical methods in the field. Metals were determined by ICP/MS after acidification of the samples.
Lignin Phenols Lignin phenols are being used in this study to examine seagrass history in selected sites in Florida Bay. In brief, fine sediment or seagrass fragments are first soxhiet extracted (methylene chloride), dried, and a weighed amount (usually @ 0.5 g) placed into monel mini bombs under an O2-free atmosphere with CuO, Fe(NH4)2(SO4)2-6H2O, and deaerated 8% NaOH. Four mini bombs at a time are placed inside a larger bomb and reacted for 3hr. 20 min. at a temperature of 170 deg C. After the reaction is complete, the bombs are quenched under running tap water and the contents of the bombs are rinsed into separate 250 ml plastic centrifuge cones with 1M NaOH. The free lignin phenols are present in the dissolved phase. The cones are centrifuged and the solutions decanted into glass round bottom flasks. The residue in the cones are washed and centrifuged twice with the 1M NaOH, which is then added to the round bottom flasks. The solutions in the round bottom flasks are then acidified to pH <2 with 6M HCl to protonate the lignin phenols. The solutions are then liquid/liquid extracted with diethyl ether (4 times) and the ether phase containing the lignin phenols isolated using a separatory funnel. The diethyl ether is dried with anhydrous Na2SO4, and blown to dryness in a small glass vial under a stream of N2. The vials are stored frozen until analysis. For analysis, the samples are redissolved in 50 ml of pyridine and derivatized with BSTFA. Samples are quantified by gas chromatography using Turbochrom software, with final confirmation of peak identities by gas chromatography/mass spectrometry using authentic standards.
U.S. Department of the Interior, U.S. Geological Survey, Center for
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