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The overall goal of this sediment biogeochemical experiment is to make realistic measurements of solute and gas exchange between the sediment surface and overlying water, with the purpose of refining aquatic mass balances and informing biogeochemical modeling efforts. This method could help answer key questions in aquatic biogeochemistry, such as how important our sediments as sinks and sources of nutrient elements, and oxygen. It is optimized for the measurement of aquatic denitrification, a major sink in the aquatic nitrogen cycle.

The main advantage of this technique is that it is highly efficient, and allows us to have considerable spatial and temporal coverage of sediment biogeochemical processes. Begin with noting the location of the sampling site by GPS. Then determine the bottom water oxygen, temperature, and salinity using a YSI water quality sonde.

Lower it to about one meter above the sediment and take a reading. Next, take readings with a PAR meter and lowering frame at the surface and at the bottom to estimate the light attenuation of the water. Now, prepare the corer.

For deep water studies, use a 6.35 centimeter inner diameter core, and for sediments with benthic microalgae or large animal populations, use a 10 centimeter inner diameter core. Now, deploy the corer over the side of the boat and lower it slowly to minimize disturbances upon penetrating the sediment. After retrieving the box corer, insert core tubes into the box.

Insert a butyl stopper to the top of each core tube, then attach an acrylic plate with an O ring to each bottom. For a pole corer, place the acrylic bottom plate in the core liner before removing the core from the corer. After coring, examine the flux core and the core box for visible disturbances or excessive resuspension.

For flux experiments, the ideal sediment-water balance within the core is 15 centimeters of each, but in coarse or highly-compacted sediments, collecting less sediment is acceptable. However, if the rates of oxygen depletion are excessive, shift the balance toward more water. Store the cores in an insulated water cooler flooded with ambient water from the site to maintain in situ temperatures.

Ensure that the cooler remains upright. Next, collect bottom water from the coring locations into 20 liter carboys using a diaphragm pump. Generally, fill two or three carboys, and collect from multiple sites if the water chemistry is anticipated to change between sites.

If a pump is unavailable in shallow, unstratified water, a carboy may be filled manually. Now, quickly transport the aerobic cores to the incubation facility before they become anoxic. Once at the facility, inspect the cores and discard any that were disturbed during transport.

Then place the cores, including the water column blank, in a bath incubator, and set the temperature to the measured bottom water temperature. Fill the incubator with the collected bottom water, completely submerging the sediment cores. A blank core, a core without any sediment, is also filled with bottom water.

Also, add bottom water to five liter carboys with spigots that will be used to dispense replacement water. For aeration, use a small T-bubbler made from 1/2 inch PVC pipe and a three-way coupler. Cover the 1/8 inch tube with a PVC pipe, and draining the water and providing circulation and aeration.

Now, let the cores equilibrate for at least two hours to overnight in the dark before sampling. Once the cores have equilibrated, attach spinning tops to the cores. With the sampling valve open, gently sweep away any air bubbles under the spinning top and seal the core from the tank water.

Next, elevate a replacement water carboy 30 to 40 centimeters above the incubation cores, then start drain lines from the carboy, and once they are flowing, attach the lines to the core tops. Then close the valves. Now, turn on the central stirring turntable and adjust the rotation speed to mix the water column without resuspending the sediment.

40 revolutions per minute usually works. After about five minutes, open the replacement water valve and the sample valve. Then, attach a short piece of tubing to the sample valve using a lure lock.

Set this sampling tube into a seven milliliter glass sample tube and overflow it with water. Then add 10 microliters of mercuric chloride to the sample and cap the tube. For solute sampling, attach a 20 milliliter syringe barrel to the sample valve and open the replacement water valve.

Then attach a plunger and a filter disc to the loaded syringe, and filter the sample into collection vials. The time course of sampling in the dark typically involves four sampling periods at half-hour to two-hour or longer intervals, depending on the rate of oxygen uptake. Using calibrated optodes, keep track of the oxygen levels in the sampled water.

In general, depletion of 25%of the oxygen allows measurement of nutrient concentration changes. Longer sampling is possible, but do not sample after 50%oxygen depletion. If the sediments are from shallow, illuminated environments, then after the fourth sample, turn on the lights and take three more samples.

Store the collected samples underwater at the incubation temperature. After taking samples, measure the remaining volume of water in the core by the water column height, or by draining all the water into a graduated cylinder. Sediment flux measurements were taken near an aquaculture facility on the Choptank River.

The rapid oxygen drop was attenuated somewhat by illumination, but the rate of photosynthesis was not as high as respiration. The control core oxygen levels changed only slightly. Nitrogen concentrations were determined using ratios of nitrogen to argon with the precision to 0.02%or about 100 nanomoles per liter of nitrogen.

In darkness or light, the sediment cores increased in nitrogen content much more than the increases observed in the water blank. Fluxes of dissolved ammonium were quite high during darkness. An increase of 20 micromoles per liter was observed in one core.

Even under illumination, the cores had increasing ammonium concentrations. Both cores and the water column blank had decreasing nitrate plus nitrite concentration over time, with lower rates of decrease during illumination. Once mastered, these incubation procedures can be carried out efficiently by two scientists, typically over the course of six to eight hours, depending on sediment respiration rates.

When attempting this procedure, it is important to remember to collect an intact sediment water interface, and to maintain a constant temperature similar to that observed in the field. After completion of the incubation, these cores can be used for sampling of the chlorophyll content, grain size, poor water chemistry, and mass-specific reactivity of the sediment. After it's development, the use of nitrogen to argon gas ratios to measure denitirification provided an alternative approach for the measurement of denitrification, and net dinitrogen fluxes.

Scientist have applied this gas ratio technique to measurement of denitrification of wetlands, lakes, reservoirs, estuaries, and shallow coastal sediments, providing a useful alternative to the isotope methodology.