William Orem Unpublished material http://sofia.usgs.gov/exchange/orem/oremsed.html The data set contains the sample ID, depth (cm), sediment size, fine sediment fraction (<60m), total C, organic C, total N, total P, C/N, C/P, and N/P. This project is examining (1) sources of nutrients, sulfur, and carbon to wetlands of south Florida, (2) the important role of chemical and biological processes to the wetland sediments (biogeochemical processes) in the cycling of these elements, and (3) the ultimate fate (i.e. sinks) of these elements in the ecosystem. The focus on nutrients and carbon reflects the problem of eutrophication in the northern Everglades, where excess phosphorus from agricultural runoff has dramatically altered the biology of the ecosystem. Results will be used by land and water managers to predict the fate of nutrients (especially phosphorus) in contaminated areas of the Everglades, and to evaluate the long-term effectiveness of buffer wetlands being constructed as nutrient removal areas. Studies of sulfur in the ecosystem are important for understanding the processes involved in mercury methylation in the Everglades. Methyl mercury (a potent neurotoxin) poses a severe health risk to organisms in the south Florida ecosystem and to humans. Sediment studies conducted by this project will also be used to construct a geochemical history of the ecosystem. An understanding of past changes in the geochemical environment of south Florida provides land and water managers with baseline information on what water quality goals for the ecosystem should be, and on how the ecosystem has responded to past environmental change and will likely respond to the changes that will accompany restoration. 19960101 19961231 ground condition complete none planned -80.75 -80.5 25.25 25 none sediment size radionuclides radioactive isotopes fine sediment fraction piston core geology peat deposits geochemistry chemistry phosphorus nitrogen carbon none Pass Key Bob Allen Key Whipray Basin Russell Key Florida Bay Monroe County Central Everglades U.S Department of Commerce, 1987, Codes for the identification of the States, the District of Columbia and the outlying areas of the United States, and associated areas (Federal Information Processing Standard 5-2): Washington, D. C., NIST Florida none None. Acknowledgement of the U.S. Geological Survey would be appreciated for products derived from these data. William Orem U.S. Geological Survey mailing address

956 National Center

Reston VA 20192 USA 703 648-6273 703 648-6419 borem@usgs.gov http://sofia.usgs.gov/workshops/waterquality/ligninphenol/fbmap.gif USGS Sediment Coring Sites in Florida Bay GIF Other USGS project personnel include Anne L. Bates, Cheryl Hedgman, Margo Corum, Ann Boylan, Robert Zielinski, Carol Kendall, Cecily Chang, J. Langston, and Sharon Fitzgerald. Outside collaborators include the South Florida Water Management District (SFWMD), the Florida Department of Environmental Protection (FDEP), Everglades Natioanl Park (NPS) and Florida Atlantic University (FAU). MS Excel Bates, Annie L. >Spiker, Elliott C. >Holmes, Charles W. 1998 Chemical Geology 146 (3-4) Amsterdam, Netherlands Elsevier Orem, William H. >Lerch, Harry E. >Rawlik, Peter 1997 USGS Open-File Reports OFR 97-454 St. Petersburg, FL U.S. Geological Survey http://sofia.usgs.gov/publications/ofr/97-454 Orem, W. H. >Holmes, C. W. >Kendall. C. >Lerch, H. E. >Bates, A. L. >Silva, S. R. >Boylan, A. >Corum, M. >Marot, M. >Hedgman, C. 1999 Journal of Coastal Research v. 15 Royal Palm Beach, FL Coastal Research and Education Foundation (CERF) The entire paper is available from the Journal of Coastal Research website at http://www.cerf-jcr-org. Journal membership is required for download. http://sofia.usgs.gov/publications/papers/geochem_flbaysed not applicable not available The positional accuracy is determined by the average of two GPS readings for each sample site. The sites are mainly accessed by helicopter. One GPS receiver is taken to the site and the other remains in the helicopter. Piston cores of sediments from various sites in Florida Bay (Pass Key, Russell Key, Bob Allen Keys, and Whipray Basin) and from nearby areas of lower Taylor Slough were returned to shore facilities (NOAA-NURC, Key Largo) for extrusion and sectioning. Cores were extruded vertically because of the generally "soupy" nature of the sediments. Cores were sectioned into 2 cm, 5 cm, or 10 cm intervals in the field, placed in ziplock plastic bags (double bagged), and refrigerated until return to lab facilities in Reston, VA. In the lab, each section was homogenized and about half of each core was wet sieved. The other half was frozen as an archive for future analyses. Brass sieves (10 mesh and 60 mesh) were used in the wet sieving. The sieves were coupled together over a large glass beaker and the sample was poured onto the 10 mesh sieve. The sample was then sequentially washed through the sieves with deionized/distilled water. Sieving produced three fractions: >10 mesh (>850 micrometers), 10-60 mesh (850-63 micrometers), and <60 mesh (<63 micrometers). Typically, the coarse and medium fractions contained mostly shells and fragments of seagrass, and the finest fraction contained only fine grained sediment. The finest fraction generally accounted for >90% of the total sediment weight. After sieving, each fraction was lyophilized, weighed, and stored in glass or plastic vials until chemical analysis. Analyses were carried out on the <63 micrometers fraction and on seagrass fragments collected from the >850 and 850-63 micrometers fractions. 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 (desiccator) 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 deg. 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 micrometer 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 precision (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). The squeezer was modified for work in south Florida from an original design by Jahnke (1988). In brief a piston core is taken using an acrylic core tube with 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 micrometers) is attached. The syringe filter is attached to a collection bottle or a syringe by a short 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. The fine sediment fraction (<63 micrometers), seagrass fragments from cores, living seagrass, and living mangrove were analyzed for lignin phenols using CuO oxidation slightly modified from the method of Hedges and Ertel (1982). 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 microliters 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. 1999 William Orem U.S. Geological Survey mailing address

956 National Center

Reston VA 20192 USA 703 648-6273 703 648-6419 borem@usgs.gov Roy Sonenshein U.S. Geological Survey Database Manager mailing address

9100 NW 36th Street Suite 107

Miami FL 33178 USA 305 717-5824 305 717-5801 sunshine@usgs.gov Florida Bay sediment geochemistry data The data have no implied or explicit guarantees EXCEL 1997 http://sofia.usgs.gov/exchange/orem/oremsed.html Log onto the SOFIA web site at http://sofia.usgs.gov none 20030728 Jo Anne Stapleton U.S. Geological Survey mailing address

521 National Center

Reston VA 20192 USA 703 648-4592 703 648-4614 jastapleton@usgs.gov Content Standard for Digital Geospatial Metadata FGDC-STD-001-1998