Salt marsh ecosystems serve as critical nutrient filters by removing reactive nitrogen (N) through denitrification. We examined the influence of long-term fertilization on N transformation and removal in a salt marsh tidal creek ecosystem fringing the Plum Island Sound estuary in northern Massachusetts, USA. Sediment oxygen demand was within the range of other marsh systems (1271.9 to 7855.0 μmol m−2 h−1) and was not significantly different between the fertilized and reference creek. Net N2 fluxes ranged from net N fixation of −402.7 μmol N2-N m−2 h−1 in the reference creek to net denitrification of 524.9 μmol N2-N m−2 h−1 in the fertilized creek. Net N2 flux and nitrate uptake were significantly higher in the fertilized creek, and in both creeks, net denitrification appeared to be nitrate limited. We calculated rates of dissimilatory nitrate reduction to ammonium (DNRA) and found it to be significantly higher in the fertilized creek, representing 45 and 11% of the total nitrate uptake in the fertilized and reference creeks, respectively. Additionally, there was a strong relationship between ammonium and nitrite fluxes in both creeks. These results suggest that DNRA may outcompete denitrification at high nitrate concentrations. Increased anthropogenic nutrient loading may therefore have a detrimental effect on the N removal capacity of salt marsh ecosystems. (From: Vieillard and Fulweiler (2012) Marine Ecology Progress Series 147: 11-22. DOI:10.3354/meps10013).
Triplicate cores were collected on 3 occasions: August 2010, June 2011, and August 2011. Samples of the tidal creek bed were taken by hand using clear polyvinyl chloride cores (5 cm diameter × 32 cm tall). The cores were inserted directly into the creek bed sediment at low tide to a depth of ~15 cm, while keeping the sediment surface undisturbed. The cores were then stored in coolers to keep them cool and dark. In the same creeks, water was collected at high tide, filtered to 0.2 μm, and stored in 20 l carboys. Both cores and filtered site water were transported back to Boston University for incubation, where they were placed in a water bath in an environmental chamber set to ambient creek water temperature.
Cores were left in the water bath overnight with air gently bubbling the overlying water. In the morning, this water was carefully siphoned out and replaced with the filtered site water. The cores were filled to the top and then fit with gas-tight lids without any air headspace or bubbles. A magnetic stir bar (~30 revo- lutions min−1) on the core tops allowed for enough water movement to prevent stratification within the core but not enough to disturb the sediment surface. Cores were incubated in the dark. Replicate water samples were collected in gas-tight 12 ml Exetainer vials (Labco) for later analysis of dissolved N2 and Ar. These samples were taken at 5 time points until the dissolved oxygen in the cores dropped at least 2 mg l−1 (62.5 μmol O2 l−1, typically 6 to 8 h). Dissolved oxygen concentrations were measured using a luminescent dissolved oxygen meter (Hach) at each time point. Once the desired drop in oxygen was attained, final samples were taken, and the incubation was stopped before the dissolved oxygen level reached 2 mg l−1 (hypoxia). The caps were then removed from the cores, and the overlying water was gently bubbled with air overnight. The next morning, the water was carefully siphoned off and replaced with filtered site water. Again, we capped the cores with a gas-tight lid and no air headspace for the second incubation for nutrient fluxes. Water samples for nutrient analysis were also collected at 5 time points until the oxygen dropped by at least 2 mg l−1. Samples for NO3−, NO2−, NH4+, and dissolved inorganic phosphorus (DIP) analysis were filtered through a 0.7 μm glass fiber filter (Whatman GF/F), collected in 30 ml acid washed, polypropylene bottles that had been leached with Milli-Q water, and stored frozen until analysis. At the end of the incubations we removed sub-samples of sediment from each core (down to 1 cm in 0.5 cm increments) for sediment density, porosity, and chlorophyll a (chl a) analysis. These samples were also frozen prior to analysis.
Frozen sediment samples were thawed, sonicated, and extracted in 25 ml of 90% acetone overnight (Dalsgaard et al. 2000). Extracted samples were then centrifuged, and 2 ml aliquots were extracted and analyzed for chl a and pheophytin fluorescence (Tril- ogy Fluorometer, Turner Designs). Sediment density and porosity were determined by drying a known volume of sediment from the top portion of each core (0 to 1 cm, in 0.5 cm increments) at ~60°C until a constant weight was achieved. Sediment density was then measured by water displacement, and porosity was also calculated (Dalsgaard et al. 2000).
(From: Vieillard and Fulweiler (2012) Marine Ecology Progress Series 147: 11-22. DOI:10.3354/meps10013).
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