1,396 research outputs found
Sediment Porosity, Density, %C, %N, and C:N ratios in Waquoit Bay, Massachusetts (USA)
No full textOn 7 occasions over the course of 3 years (2011-2013) we conducted hypoxic static core incubations on sediments and water collected in Waquoit Bay Massachusetts (USA) from four stations: Childs River Estuary, Metoxit Point, South Basin, & Sage Lot Pond. The goal of this study was quantify sediment metabolism under water column hypoxia in a shallow, temperate estuarine system. As part of that study we analyzed samples from incubated cores to determine sediment density, porosity, percent carbon (%C), percent nitrogen (%N), and molar carbon to nitrogen (C:N) ratios. These samples were collected using plastic (polycarbonate) syringe sub-cores (60 mL) that we sectioned up to 4 cm in 1 cm sub-samples. We stored the sediments in plastic centrifuge tubes (50 mL) in a freezer until they were analyzed. All sampling materials were acid washed and ethanol rinsed prior to sediment collection. We used standard processing and analysis protocols to determine sediment porosity, density (Nielsen et al. 2000) and %C %N (Zimmermann et al. 1997). Percent C and N samples were determined using an elemental analyzer at the Boston University Stable Isotopes Laboratory. For more information please see articles where these data are published (Foster & Fulweiler 2014, Foster & Fulweiler 2019). Email questions and comments to: [email protected] StationsCRE = Childs River Estuary (41° 34.805’ N 70°31.826’ W, 1.2 m deep, bottom water salinity 27.3-29.7 psu) MP = Metoxit Point (41° 34.134’ N 70° 31.272’ W, 2.2 m deep, bottom water salinity 29.6-31.3 psu) SB = South Basin (41° 33.404’ N 70° 31.442’ W, 1.8 m deep, bottom water salinity 30.6-31.3 psu)SLP = Sage Lot Pond (41° 33.270’ N 70° 30.584’ W, 1.2 m deep, bottom water salinity 28.9-30.4 psu)UnitsDepth Range = cmDensity = g/mLPorosity = sediment pore space to total volume ratioCarbon = percent carbon of total massNitrogen = percent nitrogen of total massC:N = carbon to nitrogen molar ratioAbbreviations & SymbolsDate = dd (day) - month - yy (year)Stn = Stationm.i. = measurement issuen.m. = not measuredAcknowledgmentsThere are numerous people who contributed to this project. We would like to thank the Waquoit Bay National Estuarine Research Reserve (WBNERR) for their continued multi-year support of our research. All sediment samples for this study were collected using WBNERR boats. We are particularly grateful to the following WBNERR employees who assisted with the fieldwork: MK Fox, A Lescher, J Mora, C Weidman. We would also like to thank several Fulweiler Lab members and Boston University Marine Program (BUMP) students for their assistance with fieldwork and sediment sub-coring in the lab: S Andrews, A Banks, S Buckley, K Czapla, S Donovan, D Forest, E Heiss, J Luthringer, M McCarthy, S Newell, MK Rogener, R Schweiker, K Yoshimura. In addition, S Donovan, S Duan, E Greenberg, R Lauto, and D Lewellyn, helped with sediment processing and density/porosity analysis. And R Michener in Boston University’s Stable Isotopes Laboratory analyzed sediment percent carbon and nitrogen. We also thank Boston University Earth and Environment Department for use of their facilities and their general academic and logistical research support.ReferencesFoster SQ, and RW Fulweiler. 2019. Estuarine sediments exhibit dynamic and variable biogeochemical responses to hypoxia. Journal of Geophysical Research: Biogeosciences, 124. https://doi.org/10.1029/2018JG004663Foster SQ and RW Fulweiler. 2014. Spatial and historic variability of benthic nitrogen cycling in an anthropogenically impacted estuary. Frontiers in Marine Science 1. https://doi.org/10.3389/fmars.2014.00056.Nielsen LP, V Brotas, P Viaroli, G Underwood, DB Nedwell, K Sundback, S Rysgaard, et al. 2000. Protocol handbook for NICE - Nitrogen Cycling in Estuaries: A Project under the EU reserach programme: Marine Science and Technology (MAST III). Edited by T Dalsgaard. National Environmental Research Institute, Silkborg, Denmark.Zimmermann CF, CW Keefe, and J Bashe. 1997. Determination of carbon and nitrogen in sediments and particulates of estuarine/coastal waters using elemental analysis. US Environmental Protection Agency, Method 440: 9.</div
Nutrient and dissolved gas fluxes across the sediment-water interface under normoxic conditions in Waquoit Bay, Massachusetts (USA)
No full textOn 7 occasions over the course of 3 years (2011-2013) we conducted hypoxic static core incubations on sediments and water collected in Waquoit Bay Massachusetts (USA) from four stations: Childs River Estuary, Metoxit Point, South Basin, & Sage Lot Pond. The goal of this study was quantify sediment metabolism under water column hypoxia in a shallow, temperate estuarine system. As part of that study, we compared the hypoxic flux rates to normoxic flux rates. Here we provide a comprehensive dataset of dissolved nutrient (i.e., inorganic nitrogen, phosphorus, and silica) and gas (i.e., di-nitrogen, nitrous oxide, and methane) fluxes across the sediment-water interface measured from static core incubations under normoxic conditions. Note that normoxic fluxes for O2, NH4+, PO43-, N2O, N2, are published (Foster & Fulweiler 2014; Foster & Fulweiler 2016), as are the normoxic fluxes for DSi & CH4 (Foster & Fulweiler 2019). We collected triplicate or quadruplicate sediment cores in PVC tubes (10 cm inner diameter, 30 cm height) from the side of a boat using a pole corer equipped with a one-way valve (Fulweiler et al. 2010; Foster & Fulweiler 2014). We also collected in situ bottom water from each site and filtered onboard (nominally to 0.2μm). We then transported cores and water back to our Boston University laboratory and placed them in a water-bath inside an environmental chamber set to ambient bottom water temperatures. We conducted static core incubations in the dark to determine fluxes across the sediment-water interface (e.g., Banta et al. 1995; Giblin et al. 1997; Fields et al. 2014). Core lids were equipped with magnetic stir bars (Dornblaser et al. 1989) which provided gentle (45 rpm) mixing of the overlying water with minimal suspension of sediments (Hopkinson et al. 2001; Renaud et al. 2008; Heiss et al. 2012). For each time point we collected water samples from the cores through an outflow port and water was replaced simultaneously through an inflow port to balance the volume and minimize atmospheric exchange. Dissolved inorganic nutrient concentrations were determined with high-resolution digital colorimetry on a SEAL Auto Analyzer 3 with segmented flow injection using standard techniques and chemical analyses (Solorzano 1969; Johnson & Petty 1983; Grasshoff et al. 1999). We directly measured N2 on a quadrupole membrane inlet mass spectrometer (MIMS) using the N2/Ar technique developed by Kana et al. (1994). On four occasions we also collected additional duplicate water samples for the analysis of dissolved greenhouse gases, N2O and CH4. We directly measured dissolved N2O and CH4 using a headspace equilibration technique (Kling et al. 1991; Foster & Fulweiler 2016). We analyzed the vial headspace (after water sample – headspace equilibration) using a gas chromatograph (Shimadzu GC-2014) equipped with a flame ionization detector (FID, for CH4) and an electron capture detector (ECD, for N2O). We measured oxygen concentrations using an optical Luminescent Dissolved Oxygen sensor (Hach LDO 101). Note that on one occasion (6 Aug 2012), absolute N2O flux rates were 1-3 orders of magnitude greater than on the other dates and were significant outliers in the dataset (Foster & Fulweiler 2016). These data are designated with a star (*). In addition, on 2 dates in 2012 there was an issue with the instrument analysis of N2, therefore they are designated as having a measurement issue (m.i.) and were not able to be used in our analyses. Nutrient and greenhouse gas parameters were not measured (n.m.) prior to 2012. In a few instances flux rates we could not determine (c.n.d.) flux rates because there was not predictable linear relationship between concentration change and time.Please email with questions: [email protected] StationsCRE = Childs River Estuary (41° 34.805’ N 70°31.826’ W, 1.2 m deep, bottom water salinity 27.3-29.7 psu) MP = Metoxit Point (41° 34.134’ N 70° 31.272’ W, 2.2 m deep, bottom water salinity 29.6-31.3 psu) SB = South Basin (41° 33.404’ N 70° 31.442’ W, 1.8 m deep, bottom water salinity 30.6-31.3 psu)SLP = Sage Lot Pond (41° 33.270’ N 70° 30.584’ W, 1.2 m deep, bottom water salinity 28.9-30.4 psu)Units Incubation Temperature = degrees CelsiusO2 uptake = di-oxygen per mole O2 (micromoles per meter squared per hour)NH4+ Flux = ammonium (micromoles per meter squared per hour)DSi Flux = silica (micromoles per meter squared per hour)PO43- Flux = phosphate (micromoles per meter squared per hour)N2-N Flux = di-nitrogen gas per mol N (micromoles per meter squared per hour)N2O Flux = nitrous oxide (nanomoles per meter squared per hour)CH4 Flux = methane (nanomoles per meter squared per hour)Abbreviations & SymbolsDate = dd (day) - month - yy (year)* = outlier valuem.i. = measurement issuen.m. = not measuredc.n.d. = could not determineAcknowledgmentsThere are numerous people who contributed to this project. We would like to thank the Waquoit Bay National Estuarine Research Reserve (WBNERR) for their continued multi-year support of our research. All water and sediment samples for this study were collected using WBNERR boats. We are particularly grateful to the following WBNERR employees who assisted with the fieldwork: MK Fox, A Lescher, J Mora, C Weidman. We would also like to thank several Fulweiler Lab members and Boston University Marine Program (BUMP) students for their assistance with fieldwork and the laboratory-based core incubation experiments: S Andrews, A Banks, S Buckley, K Czapla, S Donovan, D Forest, E Heiss, J Luthringer, M McCarthy, S. Newell, MK Rogener, R Schweiker, K Yoshimura. MK Rogener and E Heiss also helped analyze samples for N2/Ar concentrations on the Membrane Inlet Mass Spectrometer (MIMS). K Czapla and A Al-Haj conducted analyses for nutrient concentrations using a SEAL auto-analyzer. We also thank Boston University Earth and Environment Department for use of their facilities and their general academic and logistical research support. ReferencesBanta GT, AE Giblin, JE Hobbie, and J Tucker. 1995. Benthic respiration and nitrogen release in Buzzards Bay, Massachusetts. Journal of Marine Research 53: 107–135.Dornblaser MM, J Tucker, GT Banta, KH Foreman, MC O'Brien, and AE Giblin. 1989. Obtaining undisturbed sediment cores for biogeochemical process studies using SCUBA. In, eds. M. A. Lang and W. C. Jaap, 97–104. Costa Mesa, CA, USA.Fields L, SW Nixon, C Oviatt, and RW Fulweiler. 2014. Benthic metabolism and nutrient regeneration in hydrographically different regions on the inner continental shelf of Southern New England. Estuarine, Coastal and Shelf Science 148. Academic Press: 14–26.Foster SQ, and RW Fulweiler. 2019. Estuarine sediments exhibit dynamic and variable biogeochemical responses to hypoxia. Journal of Geophysical Research: Biogeosciences, 124. https://doi.org/10.1029/2018JG004663 Foster SQ, and RW Fulweiler. 2016. Sediment nitrous oxide fluxes are dominated by uptake in a temperate estuary. Frontiers in Marine Science 3. Frontiers: 40. https://doi.org/10.3389/fmars.2016.00040Foster SQ and RW Fulweiler. 2014. Spatial and historic variability of benthic nitrogen cycling in an anthropogenically impacted estuary. Frontiers in Marine Science 1. https://doi.org/10.3389/fmars.2014.00056.Fulweiler RW, SW Nixon, and BA Buckley. 2010. Spatial and temporal variability of benthic oxygen demand and nutrient regeneration in an anthropogenically impacted New England estuary. Estuaries and Coasts 33: 1377–1390. https://doi.org/10.1007/s12237-009-9260-y.Giblin AE, CS Hopkinson, and J Tucker. 1997. Benthic metabolism and nutrient cycling in Boston Harbor, Massachusetts. Estuaries 20: 346–364.Grasshoff K, K Kremling, and M Ehrhardt. 1999. Determination of Nutrients. In Methods of Seawater Analysis, eds. K. Grasshoff, K. Kremling, and M. Ehrhardt, 3rd ed., 159–226. Weinheim, Germany: Wiley-VCH, Verlag GmbH, D-69469.Heiss EM, L Fields, and RW Fulweiler. 2012. Directly measured net denitrification rates in offshore New England sediments. Continental Shelf Research 45. Pergamon: 78–86.Hopkinson CS, AE Giblin, and J Tucker. 2001. Benthic metabolism and nutrient regeneration on the continental shelf of Eastern Massachusetts, USA. Marine Ecology Progress Series 224: 1–19.Johnson KS, and RL Petty. 1983. Determination of nitrate and nitrite in seawater by flow injection analysis. Limnology and Oceanography 28: 1260–1266.Kling GW, GW Kipphut, and MC Miller. 1991. Arctic lakes and streams as gas conduits to the atmosphere: Implications for tundra carbon budgets. Science 251. The American Association for the Advancement of Science: 298.Renaud PE, N Morata, ML Carroll, SG Denisenko, and M Reigstad. 2008. Pelagic–benthic coupling in the western Barents Sea: processes and time scales. Deep-Sea Research Part II 55. Elsevier: 2372–2380.Solorzano L. 1969. Determination of ammonia in natural waters by the phenolypochlorite method. Limnology and Oceanography 14: 799–801.</div
Nutrient and dissolved gas fluxes across the sediment-water interface under hypoxic conditions in Waquoit Bay, Massachusetts (USA)
No full textOn 7 occasions over the course of 3 years (2011-2013) we conducted hypoxic static core incubations on sediments and water collected in Waquoit Bay Massachusetts (USA) from four stations: Childs River Estuary, Metoxit Point, South Basin, & Sage Lot Pond. The goal of this study was quantify sediment metabolism under water column hypoxia in a shallow, temperate estuarine system. Here we provide a comprehensive dataset of dissolved nutrient (i.e., inorganic nitrogen, phosphorus, and silica) and gas (i.e., di-nitrogen, nitrous oxide, and methane) fluxes across the sediment-water interface measured from static core incubations under hypoxic conditions (defined as oxygen concentrations ≤94 μM, 3 mg/L). Note that these fluxes are published (Foster & Fulweiler 2019).We collected triplicate or quadruplicate sediment cores in PVC tubes (10 cm inner diameter, 30 cm height) from the side of a boat using a pole corer equipped with a one-way valve (Fulweiler et al. 2010; Foster & Fulweiler 2014). We also collected in situ bottom water from each site and filtered onboard (nominally to 0.2μm). We then transported cores and water back to our Boston University laboratory and placed them in a water-bath inside an environmental chamber set to ambient bottom water temperatures. We conducted static core incubations in the dark to determine fluxes across the sediment-water interface (e.g., Banta et al. 1995; Giblin et al. 1997; Fields et al. 2014). Core lids were equipped with magnetic stir bars (Dornblaser et al. 1989) which provided gentle (45 rpm) mixing of the overlying water with minimal suspension of sediments (Hopkinson et al. 2001; Renaud et al. 2008; Heiss et al. 2012). In order to simulate hypoxic conditions we used the natural aerobic respiration of the sediments to consume dissolved oxygen and bring each core to hypoxia. The duration of hypoxic incubations were typically around 1 day (median 22 h, range 7.2-36 h). The median final oxygen concentrations measured in the cores at the end of the hypoxic incubation was 0.85 mg/L.Dissolved inorganic nutrient concentrations were determined with high-resolution digital colorimetry on a SEAL Auto Analyzer 3 with segmented flow injection using standard techniques and chemical analyses (Solorzano 1969; Johnson & Petty 1983; Grasshoff et al. 1999). We directly measured N2 on a quadrupole membrane inlet mass spectrometer (MIMS) using the N2/Ar technique developed by Kana et al. (1994). On four occasions we also collected additional duplicate water samples for the analysis of dissolved greenhouse gases, N2O and CH4. We directly measured dissolved N2O and CH4 using a headspace equilibration technique (Kling et al. 1991; Foster & Fulweiler 2016). We analyzed the vial headspace (after water sample – headspace equilibration) using a gas chromatograph (Shimadzu GC-2014) equipped with a flame ionization detector (FID, for CH4) and an electron capture detector (ECD, for N2O). We measured oxygen concentrations using an optical Luminescent Dissolved Oxygen sensor (Hach LDO 101). Note that on one occasion (6 Aug 2012), absolute N2O flux rates were 1-3 orders of magnitude greater than on the other dates and were significant outliers in the dataset (Foster & Fulweiler 2016). These data are designated with a star (*). In addition, on 2 dates in 2012 there was an issue with the instrument analysis of N2, therefore they are designated as having a measurement issue (m.i.) and were not able to be used in our analyses. Nutrient and greenhouse gas parameters were not measured (n.m.) prior to 2012. In a few instances flux rates we could not determine (c.n.d.) flux rates because there was not a predictable linear relationship between concentration change and time.Please email with questions: [email protected] StationsCRE = Childs River Estuary (41° 34.805’ N 70°31.826’ W, 1.2 m deep, bottom water salinity 27.3-29.7 psu) MP = Metoxit Point (41° 34.134’ N 70° 31.272’ W, 2.2 m deep, bottom water salinity 29.6-31.3 psu) SB = South Basin (41° 33.404’ N 70° 31.442’ W, 1.8 m deep, bottom water salinity 30.6-31.3 psu)SLP = Sage Lot Pond (41° 33.270’ N 70° 30.584’ W, 1.2 m deep, bottom water salinity 28.9-30.4 psu)Units Incubation Temperature = degrees CelsiusO2 uptake = di-oxygen per mole O2 (micromoles per meter squared per hour)NH4+ Flux = ammonium (micromoles per meter squared per hour)DSi Flux = dissolved silica (micromoles per meter squared per hour)PO43- Flux = phosphate (micromoles per meter squared per hour)N2-N Flux = di-nitrogen gas per mol N (micromoles per meter squared per hour)N2O Flux = nitrous oxide (nanomoles per meter squared per hour)CH4 Flux = methane (nanomoles per meter squared per hour)Abbreviations & SymbolsDate = dd (day)- month - yy (year)* = outlier value m.i. = measurement issuen.m. = not measuredc.n.d. = could not determineAcknowledgmentsThere are numerous people who contributed to this project. We would like to thank the Waquoit Bay National Estuarine Research Reserve (WBNERR) for their continued multi-year support of our research. All water and sediment samples for this study were collected using WBNERR boats. We are particularly grateful to the following WBNERR employees who assisted with the fieldwork: M.K. Fox, A. Lescher, J. Mora, C. Weidman. We would also like to thank several Fulweiler Lab members and Boston University Marine Program (BUMP) students for their assistance with fieldwork and the laboratory-based core incubation experiments: S. Andrews, A. Banks, S. Buckley, K. Czapla, S. Donovan, D. Forest, E. Heiss, J. Luthringer, M. McCarthy, S. Newell, M.K. Rogener, R. Schweiker, K. Yoshimura. M.K. Rogener and E. Heiss also helped analyze samples for N2/Ar concentrations on the Membrane Inlet Mass Spectrometer (MIMS). K. Czapla and A. Al-Haj conducted analyses for nutrient concentrations using a SEAL auto-analyzer. We also thank Boston University Earth and Environment Department for use of their facilities and their general academic and logistical research support.Text CitationsBanta GT, AE Giblin, JE Hobbie, and J Tucker. 1995. Benthic respiration and nitrogen release in Buzzards Bay, Massachusetts. Journal of Marine Research 53: 107–135.Dornblaser MM, J Tucker, GT Banta, KH Foreman, MC O'Brien, and AE Giblin. 1989. Obtaining undisturbed sediment cores for biogeochemical process studies using SCUBA. In, eds. M A Lang and W C Jaap, 97–104. Costa Mesa, CA, USA.Fields L, SW Nixon, C Oviatt, and RW Fulweiler. 2014. Benthic metabolism and nutrient regeneration in hydrographically different regions on the inner continental shelf of Southern New England. Estuarine, Coastal and Shelf Science 148. Academic Press: 14–26.Foster SQ, and RW Fulweiler. 2019. Estuarine sediments exhibit dynamic and variable biogeochemical responses to hypoxia. Journal of Geophysical Research: Biogeosciences, 124. https://doi.org/10.1029/2018JG004663Foster SQ, and RW Fulweiler. 2016. Sediment nitrous oxide fluxes are dominated by uptake in a temperate estuary. Frontiers in Marine Science 3: Article 40. https://doi.org/10.3389/fmars.2016.00040Foster SQ, and RW Fulweiler. 2014. Spatial and historic variability of benthic nitrogen cycling in an anthropogenically impacted estuary. Frontiers in Marine Science 1: Article 56. https://doi.org/10.3389/fmars.2014.00056.Fulweiler RW, SW Nixon, and BA Buckley. 2010. Spatial and temporal variability of benthic oxygen demand and nutrient regeneration in an anthropogenically impacted New England estuary. Estuaries and Coasts 33: 1377–1390. https://doi.org/10.1007/s12237-009-9260-y.Giblin AE, CS Hopkinson, and J Tucker. 1997. Benthic metabolism and nutrient cycling in Boston Harbor, Massachusetts. Estuaries 20: 346–364.Grasshoff K, K Kremling, and M Ehrhardt. 1999. Determination of Nutrients. In Methods of Seawater Analysis, eds. K Grasshoff, K Kremling, and M Ehrhardt, 3rd ed., 159–226. Weinheim, Germany: Wiley-VCH, Verlag GmbH, D-69469.Heiss EM, L Fields, and RW Fulweiler. 2012. Directly measured net denitrification rates in offshore New England sediments. Continental Shelf Research 45: 78–86.Hopkinson CS, AE Giblin, and J Tucker. 2001. Benthic metabolism and nutrient regeneration on the continental shelf of Eastern Massachusetts, USA. Marine Ecology Progress Series 224: 1–19.Johnson KS, and RL Petty. 1983. Determination of nitrate and nitrite in seawater by flow injection analysis. Limnology and Oceanography 28: 1260–1266.Kling GW, GW Kipphut, and MC Miller. 1991. Arctic lakes and streams as gas conduits to the atmosphere: Implications for tundra carbon budgets. Science 251: 298-301.Renaud PE, N Morata, ML Carroll, SG Denisenko, and M Reigstad. 2008. Pelagic–benthic coupling in the western Barents Sea: processes and time scales. Deep-Sea Research Part II 55: 2372–2380.Solorzano L. 1969. Determination of ammonia in natural waters by the phenolypochlorite method. Limnology and Oceanography 14: 799–801.</div
Sediment nitrous oxide and methane fluxes under acidification
No full textWe collected twelve sediment cores from Metoxit Point (MP) in August 2021 and from Sage Lot Pond (SLP) in September of 2021. We collected the cores using a pull corer which maintains the vertical structure of the sediment cores and preserves the delicate sediment-water interface (Foster and Fulweiler 2019). We also collected filtered bottom water (0.2 µm) in acid-washed plastic carboys. We transported the cores back to Boston University within 7 hours of collection and placed them in an environmental chamber set to 29 (±1) °C.
We conducted two separate incubations examining the effect on acidification on fluxes of nitrous oxide (N2O) and methane (CH4). We included experimental conditions representing control (pH 8.0), moderate (pH 7.3) and extreme (pH 6.3) pH conditions in Waquoit Bay (Baumann and Smith 2018). All treatments were conducted in triplicate. We divided the filtered site water into two 30-gallon acid-washed polycarbonate reservoir tanks. One reservoir tank was acidified to the appropriate pH treatment by bubbling it with 10% CO2 controlled using a Qubit pH/CO2 controller system (Qubit Systems Inc., Ontario, CA). The second tank was bubbled with compressed air to maintain a pH of 8.0.
In preparation for each incubation, we carefully siphoned off the overlying water of six sediment cores and gently replaced it with a ~4 cm headspace of treated bottom water from the corresponding station and treatment (McCarthy et al. 2013). Cores and the overlying water were capped with gas-tight plungers fitted with gas-tight PEEK inflow and outflow tubing above the sediment-water interface (overlying water volume ~314 mL). The inflow tubing was connected to a peristaltic pump (Masterflex L/S, Coleman-Palmer) with Viton (gas-impermeable) pump tubing to deliver 1.5 (± 0.2) mL min-1 of treated bottom water (overlying water residence time ~209 min; McCarthy et al. 2013). Cores were incubated in the dark under continuous flow through conditions overnight (~12 h) until sampling began the next morning (Newell et al. 2016; Li et al. 2021). During incubation, we collected water samples from inflow and outflow ports at 0, 6, 12 and 24 hours for dissolved GHG samples in 12 mL Exetainers (Labco, UK) and preserved them with 25 µL of saturated ZnCl2. Samples were kept in the refrigerator at 4 °C and analyzed within one month of sampling. After completing the first set of incubations, we repeated the process with the remaining six cores and exposed them to a moderate pH treatment.
Concentrations of dissolved N2O and CH4 were measured directly from the sample using a headspace equilibration technique and measured on a gas chromatograph (~4 mL; GC2014, Shimadzu, Japan; Foster and Fulweiler 2016). The GC was equipped with a flame ionization detector to measure CH4 and an electron capture detector to measure N2O. We estimated concentrations by comparing the area under the produced peak against a standard curve. Standard curve concentrations were determined using a linear regression of custom gas concentrations (CH4 5084 ppb; N2O: 495 ppb in N2) made by Airgas (Radnor Township, Pennsylvania, USA). Each standard curve had at least six time points and a R2 ≥ 0.995. Detection limits during sample analysis were 83.21 ppb for CH4 and 16.83 ppb for N2O.
Email questions and comments to [email protected]
Sampling Sites:
Metoxit Point = 41° 34' 8.04" N 70° 31' 17.76" W
Sage Lot Pond = 41° 34' 8.04" N 70° 30' 30.20" W
Experiment Information:
Collection_Date = Date when sediment cores were collected
Incubation_Date = Date when incubations of sediment cores began
Experiment_Type = For each station we conducted two incubations. The first incubation conducted was the Extreme treatment (pH 6.3) which included 3 control cores and 3 cores acidified to a pH of 6.3. The second experiment conducted was the moderate treatment (pH 7.3).
Treatment = The treatment each cores were exposed to:
Control = pH 8.0
pH 6.3 = Extreme
pH 7.3 = Moderate
Core_ID = Number/Letter identifying the core
Time_Point = The hour of the incubation when samples were collected
Abbreviations and Units:
Date = month - dd (day) - yy (year)
Sediment-Water Greenhouse Gas Fluxes:
N2O_Flux = nitrous oxide, nmol N2O m-2 hr-1 (nanomoles per meter squared per hour)
CH4_Flux = methane, nmol CH4 m-2 hr-1 (nanomoles per meter squared per hour)
Sediment-Water NOx Fluxes:
NOx_Flux = nitrate + nitrite, µmol m-2 hr-1 (micromoles per meter squared per hour)
Acknowledgements:
We would like to thank the Waquoit Bay National Estuarine Research Reserve (WBNERR) for their support of our research. Sediment cores for this study were collected using WBNERR boats. We are particularly grateful for Dr. Megan Tyrrell and Tonna-Marie Rogers who were instrumental in helping us carry out fieldwork and for making sure we had all the resources necessary for a successful field day. We would also like to thank members of the Fulweiler Lab, Alia Al-Haj, Amanda Vieillard, Catherine Mahoney, Kwetzpallin Mexika, Melissa Ederington Hagy, Nia Bartolucci, Paulina Azzu, and Ryan Shipley for their help with sample prep, field work and experiments.
References:
Baumann H, Smith EM (2018) Quantifying Metabolically Driven pH and Oxygen Fluctuations in US Nearshore Habitats at Diel to Interannual Time Scales. Estuaries and Coasts 41:1102–1117. https://doi.org/10.1007/s12237-017-0321-3
Foster SQ, Fulweiler RW (2019) Estuarine Sediments Exhibit Dynamic and Variable Biogeochemical Responses to Hypoxia. J Geophys Res Biogeosciences 124:737–758. https://doi.org/10.1029/2018JG004663
Foster SQ, Fulweiler RW (2016) Sediment Nitrous Oxide Fluxes Are Dominated by Uptake in a Temperate Estuary. Front Mar Sci 3:1–13. https://doi.org/10.3389/fmars.2016.00040
Li S, Twilley RR, Hou A (2021) Heterotrophic nitrogen fixation in response to nitrate loading and sediment organic matter in an emerging coastal deltaic floodplain within the Mississippi River Delta plain. Limnol Oceanogr 66:1961–1978. https://doi.org/10.1002/lno.11737
McCarthy MJ, Carini SA, Liu Z, et al (2013) Oxygen consumption in the water column and sediments of the northern Gulf of Mexico hypoxic zone. Estuar Coast Shelf Sci 123:46–53. https://doi.org/10.1016/j.ecss.2013.02.019
Newell SE, McCarthy MJ, Gardner WS, Fulweiler RW (2016) Sediment Nitrogen Fixation: a Call for Re-evaluating Coastal N Budgets. Estuaries and Coasts 39:1626–1638. https://doi.org/10.1007/s12237-016-0116-y
Funding
This work was funded by MIT SeaGrant through grants to RWF and by the NOAA Margaret A. Davidson Graduate Fellowship awarded to CIM. CIM received additional support from the Mount Holyoke Alumnae Fellowship, the Warren-McLeod Graduate Fellowship from the Boston University Marine Program, and from Boston University’s Department of Earth & Environment.</p
pH profiles in estuarine sediments exposed to acidification
No full textWe collected twelve sediment cores from two sites in Waquoit Bay (Massachusetts, USA): Metoxit Point (MP) in August 2021 and from Sage Lot Pond (SLP) in September of 2021. We collected the cores using a pull corer which maintains the vertical structure of the sediment cores and preserves the delicate sediment-water interface (Foster and Fulweiler 2019). We transported the cores back to Boston University within 7 hours of collection and placed them in an environmental chamber set to 29 (±1) °C. To better understand the impact of pH of sediment biogeochemical cycling we conducted continuous flow through incubations with treatments representing control (pH 8.0), moderate (pH 7.3) and extreme pH conditions (pH 6.3). Site water was treated with ambient air and CO2 to reach appropriate pH conditions.
Before and after each incubation we collected triplicate sediment pH profiles from each core using a dual mounted motorized micromanipulator profiler and the Sensor Trace Pro v. 3.0.6 control software (Unisense, Aarhus, Denmark). We used a Unisense pH-100 microelectrode (100 µm diameter) to collect pH measurements at 500 µM increments with a maximum profiling depth of 2 cm. We calibrated the pH probe daily with a 2-point method (Unisense, Aarhaus, Denmark) using laboratory standard buffers at pH 4.01 and pH 7.00 (Hach, USA).
Email questions and comments to [email protected]
Sampling Sites:
Metoxit Point = 41° 34’ 8.04” N 70° 31’ 17.76” W
Sage Lot Pond = 41° 34’ 8.04” N 70° 30’ 30.20” W
Profiling Information:
Depth_mm = sampling depth in millimeters
Pre_All_pH = mean pH in all cores pre incubation
Pre_All_SE = pH standard error in all cores pre incubation
Post_Ctrl_pH = mean pH in control cores post incubation
Post_Ctrl_SE = pH standard error in control cores post incubation
Post_Mod _pH = mean pH in moderate (pH 7.3) treated cores post incubation
Post_Mod_SE = pH standard error in in moderate (pH 7.3) treated cores post incubation
Post_Ext _pH = mean pH in extreme (pH 6.3) treated cores post incubation
Post_Ext_SE = pH standard error in in extreme (pH 6.3) treated cores post incubation
Acknowledgements:
We would like to thank the Waquoit Bay National Estuarine Research Reserve (WBNERR) for their support of our research. Sediment cores for this study were collected using WBNERR boats. We are particularly grateful for Dr. Megan Tyrrell and Tonna-Marie Rogers who were instrumental in helping us carry out fieldwork and for making sure we had all the resources necessary for a successful field day. We would also like to thank members of the Fulweiler Lab, Alia Al-Haj, Amanda Vieillard, Catherine Mahoney, Kwetzpallin Mexika, Melissa Ederington Hagy, Nia Bartolucci, Paulina Azzu, and Ryan Shipley for their help with sample prep, field work and experiments.
References:
Foster SQ, Fulweiler RW (2019) Estuarine Sediments Exhibit Dynamic and Variable Biogeochemical Responses to Hypoxia. J Geophys Res Biogeosciences 124:737–758. https://doi.org/10.1029/2018JG004663
Funding
This work was funded by MIT SeaGrant through grants to RWF and by the NOAA Margaret A. Davidson Graduate Fellowship awarded to CIM. CIM received additional support from the Mount Holyoke Alumnae Fellowship, the Warren-McLeod Graduate Fellowship from the Boston University Marine Program, and from Boston University’s Department of Earth & Environment.</p
Oxygen profiles in estuarine sediments exposed to acidification
No full textWe collected twelve sediment cores from two sites in Waquoit Bay (Massachusetts, USA): Metoxit Point (MP) in September 2021 and from Sage Lot Pond (SLP) in October of 2021. We collected the cores using a pull corer which maintains the vertical structure of the sediment cores and preserves the delicate sediment-water interface (Foster and Fulweiler 2019). We transported the cores back to Boston University within 7 hours of collection and placed them in an environmental chamber set to 29 (±1) °C. To better understand the impact of pH of sediment biogeochemical cycling we conducted continuous flow through incubations with treatments representing control (pH 8.0), moderate (pH 7.3) and extreme pH conditions (pH 6.3). Site water was treated with ambient air and CO2 to reach appropriate pH conditions.
We conducted triplicate sediment O2 profiles within each core before and after incubations to measure the treatment effects of pH on O2 conditions within sediments. Using a standard 100-µm O2 microelectrode (100 µm diameter) mounted on a motorized micromanipulator profiler and the Sensor Trace Pro v. 3.0.6 control software (Unisense, Aarhus, Denmark) we measured O2 concentrations at 250 µm increments down to 1 cm. All calibrations and measurements were done in the lab at ambient air temperature.
Email questions and comments to [email protected]
Sampling Sites:
Metoxit Point = 41° 34’ 8.04” N 70° 31’ 17.76” W
Sage Lot Pond = 41° 34’ 8.04” N 70° 30’ 30.20” W
Profiling Information:
Experiment Type = For each station we conducted two incubations. The first incubation conducted was the Extreme treatment (pH 6.3) which included 3 control cores and 3 cores acidified to a pH of 6.3. The second experiment conducted was the moderate treatment (pH 7.3).
Treatment = The treatment each cores were exposed to:
Control = pH 8.0
pH 6.3 = Extreme
pH 7.3 = Moderate
Incubation_Time = When profiles was collected either before incubation (Pre-Incubation) or after incubations (Post-Incubation).
Depth_mm = sampling depth in millimeters
Oxygen = oxygen concentration (µmol L-1)
SE = standard error of triplicate oxygen measurements
Group = detailed descritpion of oxygen profile
Acknowledgements:
We would like to thank the Waquoit Bay National Estuarine Research Reserve (WBNERR) for their support of our research. Sediment cores for this study were collected using WBNERR boats. We are particularly grateful for Dr. Megan Tyrrell and Tonna-Marie Rogers who were instrumental in helping us carry out fieldwork and for making sure we had all the resources necessary for a successful field day. We would also like to thank members of the Fulweiler Lab, Alia Al-Haj, Amanda Vieillard, Catherine Mahoney, Kwetzpallin Mexika, Melissa Ederington Hagy, Nia Bartolucci, Paulina Azzu, and Ryan Shipley for their help with sample prep, field work and experiments.
References
Foster SQ, Fulweiler RW (2019) Estuarine Sediments Exhibit Dynamic and Variable Biogeochemical Responses to Hypoxia. J Geophys Res Biogeosciences 124:737–758. https://doi.org/10.1029/2018JG004663
Funding
This work was funded by MIT SeaGrant through grants to RWF and by the NOAA Margaret A. Davidson Graduate Fellowship awarded to CIM. CIM received additional support from the Mount Holyoke Alumnae Fellowship, the Warren-McLeod Graduate Fellowship from the Boston University Marine Program, and from Boston University’s Department of Earth & Environment.</p
Marine macroalgae are an overlooked sink of silicon in coastal systems
No full textAcross the marine landscape, from estuaries to the open ocean, biota take up silicon (Si) as monosilicic acid and deposit it into their tissues as biogenic silica (BSi). Along the coast, vegetated ecosystems, such as salt marshes and mangroves, sequester a significant amount of Si in their tissues and likely help regulate the availability of Si in surrounding waters (Carey & Fulweiler, 2014; Elizondo et al., 2021). Si is also accumulated by sponges, euglyphid amoebae, radiolarians, silicoflagellates, and choanoflagellates, as well as a few coccolithophores, Prasinophyceae, and picocyanobacteria (Raven & Giordano, 2009; Gadd & Raven, 2010; Baines et al., 2012). The dominant driver of coastal (and open ocean) Si cycling, however, is generally thought to be diatoms. These siliceous phytoplankton require Si on a 1 : 1 molar ratio with nitrogen (N). Diatoms are responsible for 40–50% of global marine primary production (Field et al., 1998; Rousseaux & Gregg, 2013) and form the base of the marine food web in many parts of the ocean, especially coastal temperate regions (Irigoien et al., 2002)
Estimating the Aquifer’s Renewable Water to Mitigate the Challenges of Upcoming Megadrought Events
No full textIn arid and semi-arid regions of the world, the occurrence of prolonged drought events (megadroughts) associated with climate change can seriously affect the balance between water supply and demand, thereby severely increasing the susceptibility of such regions to adverse impacts. In this study, a simple framework is introduced to estimate renewable water volumes (RW) to mitigate the challenges of megadrought events by managing the groundwater resources. The framework connects a weighted annual hydrological drought index (wSPEI) to RW, based on the short time-scale precipitation volume. The proposed framework, which was in a proof-of-concept case study applied to the Neishaboor watershed in the semi-arid part of Iran, showed that developing the weighted drought index can be valuable to estimate RW. The results suggested that the wSPEI, aggregating hydrological drought index (HSPEI) with the time scale k = 5 days and the regional coefficient s = 1.3 can be used to estimate RW with reasonable accuracy (R2 = 0.73, RMSE = 11.5 mm year−1). This indicates that in the Neishaboor watershed, the best estimation of RW can be determined by precipitation volumes (or the lack thereof) falling over 5-day aggregation periods rather than by any other time scales. The accuracy of the relationship was then investigated by cross validation (leave-one-out method). According to the results, the proposed framework performed fairly well for the estimation of RW, with R2 = 0.75 and RMSE = 12.2 mm year−1 for k = 5 days. The Overall agreement between the wSPEI, the RW derived from water balance calculations, and the estimated RW by the proposed framework was also assessed for a period of 34 years. It showed that the annual RW followed closely the wSPEI, indicating a reasonable relationship between wSPEI and the annual RW. Accordingly, the proposed framework is capable to estimate the renewable water of a given watershed for different climate change scenarios.Green Open Access added to TU Delft Institutional Repository ‘You share, we take care!’ – Taverne project https://www.openaccess.nl/en/you-share-we-take-care Otherwise as indicated in the copyright section: the publisher is the copyright holder of this work and the author uses the Dutch legislation to make this work public.Water Resource
Sediment characteristics in Waquoit Bay, MA under acidification
No full textWe conducted continuous flow through incubations on sediments collected from Waquoit Bay, Massachusetts (USA) at two stations: Metoxit Point and Sage Lot Pond. The objectice of this study was to better understand the impact of moderate (pH 7.3) and extreme (pH 6.3) pH conditions on sediment nutrient and greenhouse gas fluxes. At the end of each incubation we collected samples for sediment chlorophyll-a, sediment phaeophytin, loss on ignition (LOI), percent carbon (%C), percent nitrogen (%N), molar carbon to nitrogen (C:N) ratios, and dissolved iron (DFe). We collected these samples using plastic (polycarbonate) syringe subcorers (20 mL) that we sectioned up to 4 cm in 1 cm increments. Syringes were acid washed except for those used to collect chl-a and phaeophytin. C, N, and LOI samples were stored in plastic centrifuge tubes (50 mL) in a freezer until they were analyzed. Chl-a and phaeophytin samples were stored in DI rinsed plastic centrifuge tubes (50 mL) and stored in a -80 °C freezer. DFe porewater was extracted in a glove bag under anoxic conditions, centrifuged and filtered using 0.2 µm polyethersulfone filters into DI leached and dried 2 mL polycarbonate tubes.
We followed methods for sediment % carbon, % nitrogen and C:N using the Protocol Handbook for NICE (Nitrogen Cycling in Estuaries; Dalsgaard et al. 2000). We ran these samples on an Elemental Combustion System 4010 made by Costech Analytical Technologies (Valencia, CA) with a method detection limit of 0.012 mg for C and 0.002 mg for N. LOI was determined using the dry weight after combusting samples at 600 °C for 6 hours.
We extracted and analyzed sediment chlorophyll-a and phaeophytin using techniques described by Arar and Collins (1997), Fagherazzi et al. (2014) and Ray et al. (2020).
Dissolved Fe concentrations were determined using a method described by Anschutz and Charbonnier (2021) which was adapted for micro-colorimetric assays based on a standard 96-well microplate format and read on an absorbance microplate reader (SpectraMax ABSPlus; San Jose, CA, USA).
Sampling Sites
Metoxit Point = 41° 34' 8.04" N 70° 31' 17.76" W
Sage Lot Pond = 41° 34' 8.04" N 70° 30' 30.20" W
Experiment Information
Experiment_Type = For each station we conducted two incubations. The first incubation conducted was the Extreme treatment (pH 6.3) which included 3 control cores and 3 cores acidified to a pH of 6.3. The second experiment conducted was the moderate treatment (pH 7.3).
Treatment = The acidification treatment each cores were exposed to: Control = pH 8.0
pH 6.3 = Extreme
pH 7.3 = Moderate
Core_ID: Number/Letter identifying the core
Abbreviations and Units
Collection_Date = Date of sediment collection (mm-dd-yy)
Depth = range in centimeters (cm)
Sed_Chla = sediment chlorophyll-a milligram per square meter (mg m-2)
Sed_Phaeo = sediment phaeophytin milligram per square meter (mg m-2)
C = percent carbon of total mass
N = percent nitrogen of total mass
C:N = carbon to nitrogen molar ratio
LOI = Loss on ignition
DFe = dissolved iron in porewater (µmol L-1)
Aknowledgements
We would like to thank the Waquoit Bay National Estuarine Research Reserve (WBNERR) for their support of our research. Sediment cores for this study were collected using WBNERR boats. We are particularly grateful for Dr. Megan Tyrrell and Tonna-Marie Rogers who were instrumental in helping us carry out fieldwork and for making sure we had all the resources necessary for a successful field day. We would also like to thank members of the Fulweiler Lab, Alia Al-Haj, Amanda Vieillard, Catherine Mahoney, Emily Miao, Kwetzpallin Mexika, Melissa Ederington Hagy, Nia Bartolucci, Paulina Azzu, and Ryan Shipley for their help with sample prep, field work and experiments. We also thank Dr. Cédric Fichot and Dr. Nilotpal Ghosh for help with CN sample analysis and instrumentation use.
References
Anschutz P, Charbonnier C (2021) Sampling pore water at a centimeter resolution in sandy permeable sediments of lakes, streams, and coastal zones. Limnol Oceanogr Methods 19:96–114. https://doi.org/10.1002/LOM3.10408
Arar EJ, Collins GB (1997) In Vitro Determination of Chlorophyll a and Pheophytin a in Marine and Freshwater Algae by Fluorescence. Washington, DC
Dalsgaard T, Nielsen LP, Brotas V, et al (2000) Protocol handbook for NICE - Nitrogen Cycling in Estuaries:a project under the EU research programe: Ministry of Environment and Energy National Environmental Research Institute …
Fagherazzi S, Mariotti G, Banks AT, et al (2014) The relationships among hydrodynamics, sediment distribution, and chlorophyll in a mesotidal estuary. Estuar Coast Shelf Sci 144:54–64. https://doi.org/10.1016/j.ecss.2014.04.003
Ray NE, Al-Haj AN, Fulweiler RW (2020) Sediment biogeochemistry along an oyster aquaculture chronosequence. Mar Ecol Prog Ser 646:13–27. https://doi.org/10.3354/meps13377
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Pencegahan Penyakit Infeksi Menular Melalui Edukasi PHBS Pada Masyarakat RW.02 Jungge, Kelurahan Bontoparang, Kabupaten Gowa
No full textThis service aims to provide education regarding how to prevent the emergence of infectious diseases by implementing PHBS cultural practices to the community of RW.02 Jungge. This service is carried out in the Bontoala neighborhood, Bontoparang Village, Gowa Regency. The author uses several stages in the process of service activities, including: the preparation stage, the implementation stage, and the evaluation stage. The service activity was carried out on March 9, 2022, by lecturers and students of the Faculty of Health, Patria Arta University.D uring the service process, the author found that in the preparation stage there were several gaps related to the social conditions of the RW.02 Jungge community such as social status, profession, age, educational background, economic, social, cultural, and health conditions that made the writer decide to use an educational approach. theory and practice to introduce about PHBS. Then in the second stage, namely implementation, the author and his friends did heart exercise, then continued with theoretical education regarding the importance of PHBS and how to apply PHBS using a persuasive approach, panel discussion and door to door methods. And in the third stage or evaluation stage, the writer together with other friends did a cross check by asking the RW residents one by one. 02 Jungge regarding the results and benefits of this service activity. The implementation of PKM was successfully carried out using an educational approach based on theory and practice, obtained by RW residents. O2 Jungge, Gowa Regency by knowing, understanding, and being able to prevent infectious infectious diseases by implementing PHBS.Keywords: Infectious infectious diseases, PHBS Educatio
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