The Spanish Government provided funds through the Ministerio de Educaci&#243;n as a predoctoral fellowship to C.P-L. (FPU12-00644) and research grants of the Ministerio de Economia y Competitividad: NitroPir (CGL2010-19737), Lacus (CGL2013-45348-P), Transfer (CGL2016-80124-C2-1-P). The REPLIM project (INRE - INTERREG Programme. EUUN - European Union. EFA056/15) supported the final writing of the protocol.

1. Preparation
NOTE: Begin this on the day before the measurements are taken.
<ol>
	<li>Assemble the measurement setup (Figure 1a, see the Table of Materials). 
	NOTE: To ensure a constant and high-quality power supply, the measurement device is connected to the grip via an uninterruptible power supply (UPS) that can also act as a backup. In the case of a long-duration power failure, a car battery serve as an extra power source.</li>
	<li>Start the sensor&#39;s software and apply a -0.8 V voltage to polarize the N2O microsensors. The signal shows a rapid descent and a subsequent rise, then it finally decreases until it is low and stable. 
	NOTE: The microsensor manufacturer recommends polarization at least overnight (or longer) to ensure the stability of the sensor&#39;s signal. Another recommendation is to keep the sensor polarized if measurements are planned for multiple or consecutive days18.</li>
	<li>Switch on the incubation chamber and adjust the experimental conditions (e.g., selected light off and temperature set to be similar to that expected in the field). Place a container with deionized water inside the chamber so that water is available later at the measurement temperature for calibration of the sensors. 
	NOTE: This step can be done the same day of the planned measurements, before the departure to collect the cores. For standard measurements, it is advisable to use dark conditions.</li>
	<li>Pack the field core collection materials: corer device, sampling tubes, rubber stoppers, polyvinyl chloride (PVC) taps, screwdriver, global positioning system (GPS) unit, thermometer, handheld sounder, wader, and inflatable boat (see the Table of Materials). Use a checklist to ensure that all materials are included.</li>
</ol>
2. Sediment Core Collection
<ol>
	<li>Depending on the water depth, follow 2.1.1 or 2.1.2.
	<ol>
		<li>For deep water bodies
		<ol>
			<li>Use a messenger-adapted gravity corer19 from a boat or a platform (Figure 1e).</li>
			<li>Fix the sampling tube (acrylic, &#248; 6.35 cm, length &#8805; 50 cm) to the corer with a screwdriver.</li>
			<li>Select the sampling point according to the investigation aims. Take note of the position (e.g., using GPS coordinates) and measurement depth (e.g., using a handheld sounder). If sampling from a boat, use an anchor (e.g., a bag with stones) to avoid drifting during core collection.</li>
			<li>Deploy the coring system until the sampling tube is ~1 m from the sediment. Use a rope with regular marks (e.g., intervals of 1 m) to control the depth position of the sampling equipment.</li>
			<li>Stabilize the sampling equipment for 60 s (e.g., to minimize the movement of the boat). This will ensure the correct sediment penetration and recovery of a scarcely disturbed sediment core.</li>
			<li>Release ~1 m more rope so that the sampling tube penetrates the sediment. Be aware that if the sampling tube penetrates too much, it can disturb the water/sediment interface.</li>
			<li>Release the messenger while trying to keep tension in the rope so that the corer remains fixed and in a vertical position. When the messenger impacts the corer, a small difference can be felt in the tension of the rope. At that time, close the corer to generate the vacuum that allows for recovery of the sediment core.</li>
			<li>Recover the corer by pulling the rope constantly and gently.</li>
			<li>Once the core is close to the surface but still entirely submerged (including the rubber part of the corer that ensures the vacuum), place a rubber stopper at the bottom of the sampling tube. Inspect the water/sediment interface; it should be clear and not visibly disturbed (Figure 1e). If this is not the case, discard the core, clean the tube, and repeat steps 2.1.1.4-9.</li>
			<li>Uplift the entire coring system from the water. Release the sampling tube from the corer and place a PVC cover on the top. Seal it with adhesive tape. Avoid the formation of air space.</li>
		</ol>
		</li>
		<li>For littoral habitats and shallow water bodies
		<ol>
			<li>Dress in a wader for sampling in very shallow waters (&#60;0.6 m).</li>
			<li>Use snorkeling or scuba gear for deeper sampling (up to 3 m).</li>
			<li>Select the sampling point according to the investigation aims. Take note of the position (e.g., GPS coordinates). Manually, insert the sampling tube (e.g., acrylic, &#248; 6.35 cm) into the sediment.</li>
			<li>Place a rubber stopper in the top side of the sampling tube to obtain a vacuum.</li>
			<li>Remove the core from the sediment and quickly introduce another rubber stopper at the tube bottom. 
			NOTE: It is necessary to work with the tube underwater at all times; at very shallow sites, we recommend shortening the tube down to 20 cm. Sometimes the sediment has a high water content and drains when the tube is removed from the sediment bed. In this case, it is necessary to introduce the bottom stopper without uplifting the core outside the sediment. To do this, manually immerse the stopper in the sediment around the tube and place it carefully to close the bottom of the tube.</li>
			<li>Out of the water, substitute the topside rubber stopper with a PVC cover and seal the junction with adhesive tape.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Protect the core during its transfer to the laboratory by minimizing rotations and shaking.</li>
</ol>
3. Calibration of the Nitrous Oxide (N2O) Microsensors
<ol>
	<li>Using the computer (strip chart, sensor software), check that the sensor&#39;s signal is stable and low (&#60;20 mV).</li>
	<li>Create a new file (e.g., with the date and the sampling site (130903_Redon_Lake)) to record the calibration values and sensor signals. 
	NOTE: The sensor signals are sensitive to temperature (Figure 4). Use the same temperature for the measurements and the sensor calibration. The sensor responds linearly between 0%-2.5% N2O20. Therefore, a two-point calibration is sufficient18.</li>
	<li>For the calibration value with zero nitrous oxide, read the sensor signal keeping the sensor tip submersed in N2O-free water (deionized).</li>
	<li>Calibrate with N2O water at the desired concentration. 
	&#8203;NOTE: Prepare water with a defined N2O concentration, which will slightly exceed the maximum concentration expected during incubation. We use ~25 &#181;M N2O as the calibration value. Be aware of not exceeding the maximum sensor range concentration of 500 N2O &#181;M.
	<ol>
		<li>Obtain N2O-saturated water by bubbling N2O in deionized water for a few minutes. 
		NOTE: The N2O water solubility depends on temperature and salinity21; see the table in the appendix of the sensor manual18.</li>
		<li>Dilute the N2O saturated water by adding a certain volume of saturated N2O water to a volume of deionized water. For example, at 20 &#176;C, add 0.3 mL of saturated N2O water, which has a concentration of 28.7 mM N2O, to a total of 375 mL of water to obtain a 22.9 &#181;M N2O concentration. Note that 375 mL is the total volume of the calibration chamber (Figure 1b).</li>
		<li>After gently mixing the N2O saturated water with deionized water in the calibration vessel to dilute it to the desired concentration, read the sensor signal when it is constant. This reading is the calibration value with X &#181;M N2O water. When mixing the solution, be careful not to generate bubbles, as this would eliminate N2O from the calibration solution. 
		NOTE: Be aware that the N2O in the water will slowly escape into the air; thus, the prepared calibration solution can only be used for a few minutes.</li>
	</ol>
	</li>
</ol>
4. Core Preparation and Acetylene Inhibition
<ol>
	<li>Change the PVC cover located at the top of each sediment core by another cover with a hole in the center and a hanging magnetic stirrer. Re-seal the junction with adhesive tape.</li>
	<li>Reduce the water phase of each sample to an approximate height of 12 cm (volume &#8776; 380 mL). For this, first insert a silicone tube in the central hole.Then, put the sediment core in a cylinder and push the bottom stopper to create pressure. The stopper and sediment sample go up, and the excess water passes through the tube. Collect the water in a recipient vessel. 
	NOTE: Samples with coarse granularity can be problematic during this step. Sediment particles placed between the stopper and the tube can deform the stopper and open a hole through which air bubbles can pass and disturb the sample. To avoid this problem, put the cylinder in the center of the bottom stopper and try to push with a constant force. The joint between the silicone tube used to evacuate the excess water and the PVC cover consists of a solid part (e.g., a 5 mL pipette tip without its narrowest end) inserted in the silicone tube.</li>
	<li>Perform the acetylene inhibition by bubbling with acetylene gas in the water phase of the core for approximately 10 min . Avoid resuspending the sediment. 
	NOTE: As a possible modification of the method, add a substrate (nitrate) through a concentrated liquid medium before bubbling acetylene for potential denitrification measurements (e.g., as in Figure 3b, c).</li>
</ol>
5. Denitrification (N2O accumulation measure)
<ol>
	<li>Fill all the air space in the sample with the previous leftover water. Place the sensor in the sediment core through the central hole of the topside PVC cover. The tip of the sensor should be located in the water phase above the stirrer (Figure 1c). 
	NOTE: All the joints of the acrylic sampling tube must be sealed to avoid gas and water leaks during the measurement (Figure 1a, c). In the bottom part of the tube, the rubber stopper is sufficient for this. Sealing the topside part is more difficult. The PVC cover must be tuned. It must be heated with a torch; then, when the material becomes flexible but is not scorched, the cover is placed in the tube so that its shape can be molded. After cooling, the cover needs more modifications (with the exception of the cover used to transport the samples to the laboratory in steps 2.1.1.10 or 2.1.2.6). The central hole where the sensor is inserted must be drilled. The stirrer can be held with a fishing line, which in turn is adhered with glue to the inside of the cover so that the stirrer hangs on the fishing line in the water (Figure 1c). Also, all the joints (PVC cover tube and PVC cover sensor) are sealed with adhesive tape. Place elastic adhesive tape to adjust the diameter of the sensor in order to seal the contact surface between the central hole of the PVC cover and the sensor (Figure 1c).</li>
	<li>Switch on the electromagnetic pulse circuit that is part of the stirring system. 
	NOTE: The stirring system prevents the stratification of the water phase without disturbing (resuspending) the sediment. The stirring system consists of a circuit that switches on/off the electromagnet that attracts/releases the magnetic stirrer (see the Table of Materials for a detailed description).</li>
	<li>Move the electromagnet around the external part of the acrylic tube until the stirrer moves continuously, and then fix it in place using adhesive tape (Figure 1c).</li>
	<li>Close the incubation chamber to ensure a constant temperature (e.g., variation of &#177;0.3 &#176;C).</li>
	<li>Press the record button (sensor software) to start recording the sensor signal. Readings are typically recorded every 5 min.</li>
	<li>Press the stop button at the end of the measurement period.</li>
</ol>
6. Final Measurement Steps
<ol>
	<li>Wait at least ~10 min with the sensor&#39;s tip submerged in free-N2O water (deionized) before reading the signal of the zero N2O calibration measure.</li>
	<li>Perform a final sensor calibration. For this, repeat the sensor calibration, following Section 3 but starting with step 3.3.</li>
	<li>Save the file (sensor software).</li>
</ol>
7. Denitrification Rate Calculations
<ol>
	<li>Start with the tabulated output file generated by the sensor software that contains the record of the sensor&#39;s signal in mV and &#181;M N2O, and the calibration data.</li>
	<li>Plot the sensor signal against time to visualize the N2O accumulation trend (e.g., Figure 2a).</li>
	<li>Use only the time range with a linear accumulation, excluding the initial acclimation period of the sample and a possible final saturation due to substrate limitation (e.g., Figure 2b). Create a linear model of the sensor signal (&#181;M) over time (h) . 
	Note: The slope is the denitrification rate (&#181;M N2O core-1 h-1), which, if divided by the area of the core (&#960;r2), transforms into the rate in &#181;M N2O m-2 h-1, and when multiplied by the water volume (&#960;r2h, where h is the height of the water phase and r is the inner radius of the acrylic tube, in this case 0.12 m and 0.03175 m, respectively) transforms into the rate in &#181;mol N2O m-2 h-1.</li>
</ol>

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B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Denitrification+measurements+in+aquatic+sediments+-+a+comparison+of+3+methods.">Denitrification measurements in aquatic sediments - a comparison of 3 methods.</a> Biogeochemistry. 23 (3), 147-167 (1993).</li><li>Christensen, P. B., Nielsen, L. P., Revsbech, N. P., Sorensen, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microzonation+of+denitrification+activity+in+stream+sediments+as+studied+with+a+combined+oxygen+and+nitrous-oxide+microsensor.">Microzonation of denitrification activity in stream sediments as studied with a combined oxygen and nitrous-oxide microsensor.</a> Applied and Environmental Microbiology. 55 (5), 1234-1241 (1989).</li><li>Peter, N. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Denitrification+in+sediment+determined+from+nitrogen+isotope+pairing.">Denitrification in sediment determined from nitrogen isotope pairing.</a> FEMS Microbiology Ecology. 9 (4), 357-361 (1992).</li><li>Risgaard-Petersen, N., Nielsen, L. P., Rysgaard, S., Dalsgaard, T., Meyer, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+of+the+isotope+pairing+technique+in+sediments+where+anammox+and+denitrification+coexist.">Application of the isotope pairing technique in sediments where anammox and denitrification coexist.</a> Limnology and Oceanography-Methods. 1, 63-73 (2003).</li></ol>

The authors have nothing to disclose.

The main advantages of the described method are the use of minimally disturbed sediment core samples and the continuous recording of the N2O accumulation. These allow estimation of relatively low denitrification rates that are likely similar to those occurring in situ. Nonetheless, some aspects concerning the coring, sensor performance, and potential improvements are discussed.
An apparently simple but critical step of the method is good core recovery. The sediment/water in...

Anthropogenic alteration of the nitrogen cycle is one of the most challenging problems for the Earth system1. Human activity has at least doubled the levels of reactive nitrogen available to the biosphere2. However, there remain large uncertainties regarding how the global N cycle is evaluated. A few flux estimates have been quantified with less than &#177;20% error, and many have uncertainties of &#177;50% and larger3. These uncertainties indicate the need for accurate estimations of denitrification rates across ecosystems and an understanding of the underlying mechanisms of variation. Denitrification is a microbial activity through which nitrogenous oxides, mainly nitrate and nitrite, are reduced to dinitrogen gasses, N2O and N24. The pathway is highly relevant to the biosphere availability of reactive nitrogen because it is the primary process of removal5. N2O is a greenhouse gas with a warming potential nearly 300 times that of CO2 over 100 years, and it is the current major cause of stratospheric ozone depletion due to the large quantities being emitted6,7.
In the following, we present a protocol for estimating sediment denitrification rates using cores and N2O microsensors experimentally (Figure 1). Denitrification rates are estimated using the acetylene inhibition method8,9 and measurements of the accumulation of N2O during a defined period (Figure 2 and Figure&#160;3). We demonstrate the method by applying it to mountain lake sediments. This case study highlights the performance of the method for detecting relatively low rates with minimal disturbance to the physical structure of the sediments.
Denitrification is particularly difficult to measure10. There are several alternative approaches and methods, each with advantages and disadvantages. Drawbacks to available methods include their use of expensive resources, insufficient sensitivity, and the need to modify the substrate levels or alter the physical configuration of the process using disturbed samples10. An even more fundamental challenge to measuring N2 is its elevated background levels in the environment10. The reduction of N2O to N2 is inhibited by acetylene (C2H2)8,9. Thus, denitrification can be quantified by measuring the accumulated N2O in the presence of C2H2, which is feasible due to low environmental N2O levels.
The use of C2H2 to measure denitrification rates in sediments was developed about 40 years ago11, and the incorporation of N2O sensors occurred about 10 years later12. The most widely applied acetylene-based approach is the &#34;static core&#34;. The accumulated N2O is measured during an incubation period of up to 24 h after the C2H2 is added to the headspace of the sealed sediment core10. The method described here follows this procedure with some innovations. We add the C2H2 by bubbling the gas in the water phase of the core for some minutes, and we fill all the headspace with sample water before measuring the accumulation of N2O with a microsensor. We also include a stirring system that prevents the stratification of the water without resuspending the sediment. The procedure quantifies the denitrification rate per sediment surface area (e.g., &#181;mol N2O m-2 h-1).
The high spatial and temporal variation of denitrification presents another difficulty in its accurate quantification10. Usually, N2O accumulation is measured sequentially by gas chromatography of headspace samples that are collected during the incubation. The method described provides improved monitoring of the temporal variation of the N2O accumulation, because the microsensor provides a continuous signal. The microsensor multimeter is a digital microsensor amplifier (picoammeter) that interfaces with the sensor(s) and the computer (Figure 1a). The multimeter allows several N2O microsensors to be used at the same time. For instance, up to four sediment cores from the same study site can be measured simultaneously to account for the spatial variability.
The core approach barely disturbs the sediment structure compared to some other methods (e.g., slurries). If the integrity of the sediments is altered, this leads to unrealistic denitrification rates13 that are only adequate for relative comparisons. Higher rates are always obtained with slurry methods compared to core methods14, because the latter preserves the limitation of denitrification by substrate diffusion15. Slurry measures cannot be considered representative of in situ rates16; they provide relative measures for comparisons made with the exact same procedure.
The method described is appropriate for estimating denitrification rates in any sediment type that can be cored. We particularly recommend the method for performing experimental manipulations of some of the driving factors. Examples are experiments that modify nitrate availability and temperature as needed for estimating the energy activation (Ea) of denitrification17 (Figure 2).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58553/58553fig1v2.jpg" /> 
Figure 1: Experimental setup. (a) General experimental setup to estimate sediment denitrification rates using cores and N2O microsensors. The incubation chamber ensures darkness and controlled-temperature (&#177;0.3 &#176;C) conditions. Five intact sediment cores can be processed simultaneously using their respective N2O sensors. (b) N2O sensor calibration chamber. We adapted it with rubber stoppers and syringes to mix the N2O water (see protocol step 3.4.3). There is a thermometer to control the water temperature. (c) Close-up of a sediment core sample with the sensor inserted into the central hole of the PVC cover and the joints sealed with adhesive tape. The stirrer is hanging in the water, and the electromagnet is close to it and fixed to the external part of the acrylic tube. (d) Close-up of the N2O microsensor tip protected by a metal piece. (e) A sediment core that has just been recovered. It was sampled from a boat in a deep lake; the acrylic tube with the core is still fixed to the messenger-adapted gravity corer19. See the Table of Materials for all the items needed to perform this method. <a href="https://www.jove.com/files/ftp_upload/58553/58553fig1v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="0" cellpadding="0" cellspacing="0" style="width:595px;" width="594">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="63">
			<td height="63" style="height:63px;width:227px;">Messenger-adapted gravity corer</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Reference in the manuscript. Made by Glew, J.</td>
		</tr>
		<tr height="20">
			<td height="20" style="height:20px;width:227px;">Sampling tube</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Acrylic. Dimensions: 100 cm (h) &#215; 6.35 cm (d) &#215; 6.50 cm (D). Sharpen the edge of the sampling tube that penetrates into the sediment to minimize the disturbance in the recovered sediment core sample.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:227px;">Handheld sounder</td>
			<td style="width:65px;">Plastimo</td>
			<td style="width:100px;">38074</td>
			<td style="width:203px;">Echotest II Depth Sounder.</td>
		</tr>
		<tr height="128">
			<td height="128" style="height:128px;width:227px;">Rubber stopper</td>
			<td style="width:65px;">VWR</td>
			<td style="width:100px;">DENE1012114</td>
			<td style="width:203px;">With two holes, used to mix the N2O-water in the calibration chamber. Dimensions: 20 mm (h) &#215; 14 mm (d) &#215; 18 mm (D) (3 mm hole (D)).</td>
		</tr>
		<tr height="126">
			<td height="126" style="height:126px;width:227px;">Rubber stopper</td>
			<td style="width:65px;">VWR</td>
			<td style="width:100px;">217-0125</td>
			<td style="width:203px;">To seal the bottom part of the methacrylate tube and to sample in shallow water bodies. Dimensions: 45 mm (h) &#215; 56 mm (d) &#215; 65 mm (D).</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:227px;">PVC cover</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">To seal the top side part of the acrylic tube. Dimensions: 45 mm (h) &#215; 56 mm (d) &#215; 65 mm (D). Dimensions: 65 mm (D).</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:227px;">Adhesive tape</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Waterproof. To ensure all joints (PVC cover sampling tube and PVC cover sensor) and to avoid water leaks.</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:227px;">Thermometer</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Portable and waterproof, to measure the temperature in the water overlying the sediment just after sampling the cores.</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:227px;">GPS</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">To save the location of a new sampling site or to arrive at a previous site.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:227px;">Wader</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">For littoral or shallow site samplings.</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:227px;">Boat</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">An inflatable boat is the best option for its lightness if the sampling site is not accessible by car.</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:227px;">Rope</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Rope with marks showing its length (e.g., marked with a color code to distinguish each meter).</td>
		</tr>
		<tr height="44">
			<td height="44" style="height:44px;width:227px;">N2O gas bottle and pressure reducer</td>
			<td style="width:65px;">Abell&#243; Linde</td>
			<td style="width:100px;">32768-100</td>
			<td style="width:203px;">Gas bottle reference.</td>
		</tr>
		<tr height="44">
			<td height="44" style="height:44px;width:227px;">C2H2 gas bottle and pressure reducer</td>
			<td style="width:65px;">Abell&#243; Linde</td>
			<td style="width:100px;">32468-100</td>
			<td style="width:203px;">Gas bottle reference.</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:227px;">Tube used to evacuate the excess of water</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Consists of a solid part (e.g., a 5 ml pipette tip without its narrowest end) inserted in a silicone tube.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:227px;">Nitrous Oxide Minisensor w/ Cap</td>
			<td style="width:65px;">Unisense</td>
			<td style="width:100px;">N2O-R</td>
			<td style="width:203px;">We use 4 sensors at a time.</td>
		</tr>
		<tr height="235">
			<td height="235" style="height:235px;width:227px;">Microsensor multimeter 4 Ch. 4 pA channels</td>
			<td style="width:65px;">Unisense</td>
			<td style="width:100px;">Multimeter</td>
			<td style="width:203px;">Picoammeter logged to a laptop. The standard device allows for 2 sensor picoammeter connections (e.g., N2O sensor), one pH/mV and a thermometer. We ordered a device with four picoammeter connections, allowing the use of 4 N2O sensors simultaneously.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:227px;">SensorTrace Basic 3.0 Windows software</td>
			<td style="width:65px;">Unisense</td>
			<td style="width:100px;"></td>
			<td style="width:203px;">Sensor data acquisition software.</td>
		</tr>
		<tr height="107">
			<td height="107" style="height:107px;width:227px;">Calibration Chamber incl. pump</td>
			<td style="width:65px;">Unisense</td>
			<td style="width:100px;">CAL300</td>
			<td style="width:203px;">Calibration chamber. We tuned it with rubber stoppers and syringes to mix the N2O-water without making bubbles.</td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:227px;">Incubation chamber</td>
			<td style="width:65px;">Ibercex</td>
			<td style="width:100px;">E-600-BV</td>
			<td style="width:203px;">Indispensable equipment for working at a constant temperature (&#177;0.3 &#176;C). It also allows control of the photoperiod.</td>
		</tr>
		<tr height="126">
			<td height="126" style="height:126px;width:227px;">Electric stirrer</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Part of the stirring system. It hangs in the water, overlying the sediment subject, by a fishing line that is hooked to the PVC cover.</td>
		</tr>
		<tr height="252">
			<td height="252" style="height:252px;width:227px;">Electromagnet</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Part of the stirring system. It is fixed to the outside of the acrylic tube, approximately at the same level as the stirrer. It is activated episodically (ca. 1 on-off per s) by a circuit, attracting the stirrer when it is on and releasing it when it is off, thereby generating the movement that agitates the water.</td>
		</tr>
		<tr height="126">
			<td height="126" style="height:126px;width:227px;">Electromagnetic pulse circuit</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">Part of the stirring system. It is connected by wires to the electromagnet and sends pulses of current that turn the electromagnet on and off.</td>
		</tr>
		<tr height="170">
			<td height="170" style="height:170px;width:227px;">Uninterruptible power supply (UPS)</td>
			<td style="width:65px;">-</td>
			<td style="width:100px;">-</td>
			<td style="width:203px;">It improves the quality of the electrical energy that reaches the measurement device, filtering the highs and low of the voltage, thereby ensuring a more constant and stable N2O sensor signal.</td>
		</tr>
	</tbody>
</table>

estimating sediment denitrification rates using cores and n2o microsensors

Denitrification is the primary biogeochemical process removing reactive nitrogen from the biosphere. The quantitative evaluation of this process has become particularly relevant for assessing the anthropogenic-altered global nitrogen cycle and the emission of greenhouse gases (i.e., N2O). Several methods are available for measuring denitrification, but none of them are completely satisfactory. Problems with existing methods include their insufficient sensitivity, and the need to modify the substrate levels or alter the physical configuration of the process using disturbed samples.This work describes a method for estimating sediment denitrification rates that combines coring, acetylene inhibition, and microsensor measurements of the accumulated N2O. The main advantages of this method are a low disturbance of the sediment structure and the collection of a continuous record of N2O accumulation; these enable estimates of reliable denitrification rates with minimum values up to 0.4-1 &#181;mol N2O m-2 h-1. The ability to manipulate key factors is an additional advantage for obtaining experimental insights. The protocol describes procedures for collecting the cores, calibrating the sensors, performing the acetylene inhibition, measuring the N2O accumulation, and calculating the denitrification rate. The method is appropriate for estimating denitrification rates in any aquatic system with retrievable sediment cores. If the N2O concentration is above the detection limit of the sensor, the acetylene inhibition step can be omitted to estimate the N2O emission instead of denitrification. We show how to estimate both actual and potential denitrification rates by increasing nitrate availability as well as the temperature dependence of the process. We illustrate the procedure using mountain lake sediments and discuss the advantages and weaknesses of the technique compared to other methods. This method can be modified for particular purposes; for instance, it can be combined with 15N tracers to assess nitrification and denitrification or field in situ measurements of denitrification rates.

A total of 468 denitrification rates were estimated using the protocol above in sediments from Pyrenean mountain lakes over the period 2013-2014. We show some of these results to illustrate the procedure (Figure 2 and Figure&#160;3). In general, the linear model between the N2O concentration and time has good correlation (R2 &#8805; 0.9). The slope of the relationship provides an estimate of the denitrificat...

Watch this Scientific Journal Video about Estimating Sediment Denitrification Rates Using Cores and N2O Microsensors at JoVE.com

Estimating Sediment Denitrification Rates Using Cores and N2O Microsensors

This method estimates sediment denitrification rates in sediment cores using the acetylene inhibition technique and microsensor measurements of the accumulated N2O. The protocol describes procedures for collecting the cores, calibrating the sensors, performing the acetylene inhibition, measuring the N2O accumulation, and calculating the denitrification rate.

Estimating Sediment Denitrification Rates Using Cores and N2O Microsensors

Denitrification is the primary biogeochemical process removing reactive nitrogen from the biosphere. The quantitative evaluation of this process has ...

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8.2K Views.  CREAF. This method estimates sediment denitrification rates in sediment cores using the acetylene inhibition technique and microsensor measurements of the accumulated N2O. The protocol describes procedures for collecting the cores, calibrating the sensors, performing the acetylene inhibition, measuring the N2O accumulation, and calculating the denitrification rate.

Video: Estimating Sediment Denitrification Rates Using Cores and N2O Microsensors

Transient Gene Expression in Tobacco using Gibson Assembly and the Gene Gun

In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately using complex regulatory strategies including differential promoters and/or signal sequences. Metabolic engineers and synthetic biologists interested in targeting enzymes to a particular organelle are faced with a challenge: For a protein that is to be localized to more than one organelle, the engineer must clone the same gene multiple times. This work presents a solution to this strategy: harnessing alternative splicing of mRNA. This technology takes advantage of established chloroplast and peroxisome targeting sequences and combines them into a single mRNA that is alternatively spliced. Some splice variants are sent to the chloroplast, some to the peroxisome, and some to the cytosol. Here the system is designed for multiple-organelle targeting with alternative splicing. In this work, GFP was expected to be expressed in the chloroplast, cytosol, and peroxisome by a series of rationally designed 5&#8217; mRNA tags. These tags have the potential to reduce the amount of cloning required when heterologous genes need to be expressed in multiple subcellular organelles. The constructs were designed in previous work11, and were cloned using Gibson assembly, a ligation independent cloning method that does not require restriction enzymes. The resultant plasmids were introduced into Nicotiana benthamiana epidermal leaf cells with a modified Gene Gun protocol. Finally, transformed leaves were observed with confocal microscopy.

This work describes a novel method for selectively targeting subcellular organelles in plants, assayed using the BioRad Gene Gun.

In order to target a single protein to multiple subcellular organelles, plants typically duplicate the relevant genes, and express each gene separately ...

Measuring Rates of Herbicide Metabolism in Dicot Weeds with an Excised Leaf Assay

In order to isolate and accurately determine rates of herbicide metabolism in an obligate-outcrossing dicot weed, waterhemp (Amaranthus tuberculatus), we developed an excised leaf assay combined with a vegetative cloning strategy to normalize herbicide uptake and remove translocation as contributing factors in herbicide-resistant (R) and &#8211;sensitive (S) waterhemp populations. Biokinetic analyses of organic pesticides in plants typically include the determination of uptake, translocation (delivery to the target site), metabolic fate, and interactions with the target site. Herbicide metabolism is an important parameter to measure in herbicide-resistant weeds and herbicide-tolerant crops, and is typically accomplished with whole-plant tests using radiolabeled herbicides. However, one difficulty with interpreting biokinetic parameters derived from whole-plant methods is that translocation is often affected by rates of herbicide metabolism, since polar metabolites are usually not mobile within the plant following herbicide detoxification reactions. Advantages of the protocol described in this manuscript include reproducible, accurate, and rapid determination of herbicide degradation rates in R and S populations, a substantial decrease in the amount of radiolabeled herbicide consumed, a large reduction in radiolabeled plant materials requiring further handling and disposal, and the ability to perform radiolabeled herbicide experiments in the lab or growth chamber instead of a greenhouse. As herbicide resistance continues to develop and spread in dicot weed populations worldwide, the excised leaf assay method developed and described herein will provide an invaluable technique for investigating non-target site-based resistance due to enhanced rates of herbicide metabolism and detoxification.

This manuscript describes how herbicide metabolism rates can be effectively quantified with excised leaves from a dicot weed, thereby reducing variability and removing any possible confounding effects of herbicide uptake or translocation typically observed in whole-plant assays.

In order to isolate and accurately determine rates of herbicide metabolism in an obligate-outcrossing dicot weed, waterhemp (Amaranthus tuberculatus), we ...

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

We demonstrate a method for sectioning sediment cores and extracting pore waters while maintaining oxygen-free conditions. A simple, inexpensive system is built and can be transported to a temporary work space close to field sampling site(s) to facilitate rapid analysis. Cores are extruded into a portable glove bag, where they are sectioned and each 1-3 cm thick section (depending on core diameter) is sealed into 50 ml centrifuge tubes. Pore waters are separated with centrifugation outside of the glove bag and then returned to the glove bag for separation from the sediment. These extracted pore water samples can be analyzed immediately. Immediate analyses of redox sensitive species, such as sulfide, iron speciation, and arsenic speciation indicate that oxidation of pore waters is minimal; some samples show approximately 100% of the reduced species, e.g. 100% Fe(II) with no detectable Fe(III). Both sediment and pore water samples can be preserved to maintain chemical species for further analysis upon return to the laboratory.

A protocol for sectioning sediment cores and extracting pore waters under anoxic conditions in order to permit analysis of redox sensitive species in both solids and fluids is presented.

We demonstrate a method for sectioning sediment cores and extracting pore waters while maintaining oxygen-free conditions. A simple, inexpensive system is ...

Continuous Instream Monitoring of Nutrients and Sediment in Agricultural Watersheds

Pollutant concentrations and loads in watersheds vary considerably with time and space. Accurate and timely information on the magnitude of pollutants in water resources is a prerequisite for understanding the drivers of the pollutant loads and for making informed water resource management decisions. The commonly used &#34;grab sampling&#34; method provides the concentrations of pollutants at the time of sampling (i.e., a snapshot concentration) and may under- or overpredict the pollutant concentrations and loads. Continuous monitoring of nutrients and sediment has recently received more attention due to advances in computing, sensing technology, and storage devices. This protocol demonstrates the use of sensors, sondes, and instrumentation to continuously monitor in situ nitrate, ammonium, turbidity, pH, conductivity, temperature, and dissolved oxygen (DO) and to calculate the loads from two streams (ditches) in two agricultural watersheds. With the proper calibration, maintenance, and operation of sensors and sondes, good water quality data can be obtained by overcoming challenging conditions such as fouling and debris buildup. The method can also be used in watersheds of various sizes and characterized by agricultural, forested, and/or urban land.

With the advancement of technology and the rise in end-user expectations, the need and use of higher temporal resolution data for pollutant load estimation has increased. This protocol describes a method for continuous in situ water quality monitoring to obtain higher temporal resolution data for informed water resource management decisions.

Pollutant concentrations and loads in watersheds vary considerably with time and space. Accurate and timely information on the magnitude of pollutants in ...

Preparation of DMMTAV and DMDTAV Using DMAV for Environmental Applications: Synthesis, Purification, and Confirmation

Dimethylated thioarsenicals such as dimethylmonothioarsinic acid (DMMTAV) and dimethyldithioarsinic acid (DMDTAV), which are produced by the metabolic pathway of dimethylarsinic acid (DMAV) thiolation, have been recently found in the environment as well as human organs. DMMTAV and DMDTAV can be quantified to determine the ecological effects of dimethylated thioarsenicals and their stability in environmental media. The synthesis method for these compounds is unstandardized, making replicating previous studies challenging. Furthermore, there is a lack of information about storage techniques, including storage of compounds without species transformation. Moreover, because only limited information about synthesis methods is available, there may be experimental difficulties in synthesizing standard chemicals and performing quantitative analysis. The protocol presented herein provides a practically modified synthesis method for the dimethylated thioarsenicals, DMMTAV and DMDTAV, and will help in the quantification of species separation analysis using high performance liquid chromatography in conjunction with inductively coupled plasma mass spectrometry (HPLC-ICP-MS). The experimental steps of this procedure were modified by focusing on the preparation of chemical reagents, filtration methods, and storage.

Preparation of DMMTAV and DMDTAV Using DMAV for Environmental Applications: Synthesis, Purification, and Confirmation

This article presents modified experimental protocols for dimethylmonothioarsinic acid (DMMTAV) and dimethyldithioarsinic acid (DMDTAV) synthesis, inducing dimethylarsinic acid (DMAV) thiolation through mixing of DMAV, Na2S, and H2SO4. The modified protocol provides an experimental guideline, thereby overcoming limitations of the synthesis steps that could have caused experimental failures in quantitative analysis.

Dimethylated thioarsenicals such as dimethylmonothioarsinic acid (DMMTAV) and dimethyldithioarsinic acid (DMDTAV), which are produced by the metabolic ...

Sampling, Sorting, and Characterizing Microplastics in Aquatic Environments with High Suspended Sediment Loads and Large Floating Debris

The ubiquitous presence of plastic debris in the ocean is widely recognized by the public, scientific communities, and government agencies. However, only recently have microplastics in freshwater systems, such as rivers and lakes, been quantified. Microplastic sampling at the surface usually consists of deploying drift nets behind either a stationary or moving boat, which limits the sampling to environments with low levels of suspended sediments and floating or submerged debris. Previous studies that employed drift nets to collect microplastic debris typically used nets with &#8805;300 &#181;m mesh size, allowing plastic debris (particles and fibers) below this size to pass through the net and elude quantification. The protocol detailed here enables: 1) sample collection in environments with high suspended loads and floating or submerged debris and 2) the capture and quantification of microplastic particles and fibers &lt;300 &#181;m. Water samples were collected using a peristaltic pump in low-density polyethylene (PE) containers to be stored before filtering and analysis in the lab. Filtration was done with a custom-made microplastic filtration device containing detachable union joints that housed nylon mesh sieves and mixed cellulose ester membrane filters. Mesh sieves and membrane filters were examined with a stereomicroscope to quantify and separate microplastic particulates and fibers. These materials were then examined using a micro-attenuated total reflectance Fourier transform infrared spectrometer (micro ATR-FTIR) to determine microplastic polymer type. Recovery was measured by spiking samples using blue PE particulates and green nylon fibers; percent recovery was determined to be 100% for particulates and 92% for fibers. This protocol will guide similar studies on microplastics in high velocity rivers with high concentrations of sediment. With simple modifications to the peristaltic pump and filtration device, users can collect and analyze various sample volumes and particulate sizes.

Most microplastic research to date has occurred in marine systems where suspended solid levels are relatively low. Focus is now shifting to freshwater systems, which may feature high sediment loads and floating debris. This protocol addresses collecting and analyzing microplastic samples from aquatic environments that contain high suspended solid loads.

The ubiquitous presence of plastic debris in the ocean is widely recognized by the public, scientific communities, and government agencies. However, only ...

Laboratory and Field Protocol for Estimating Sheet Erosion Rates from Dendrogeomorphology

Sheet erosion is among the crucial drivers of soil degradation. Erosion is controlled by environmental factors and human activities, which often lead to severe environmental impacts. The understanding of sheet erosion is, consequently, a worldwide issue with implications for both environment and economies. However, the knowledge on how erosion evolves in space and time is still limited, as well as its effects on the environment. Below, we explain a new dendrogeomorphological protocol for deriving eroded soil thickness (Ex) by acquiring accurate microtopographic data using both terrestrial laser scanning (TLS) and microtopographic profile gauges. Additionally, standard dendrogeomorphic procedures, dependent on anatomical variations in root rings, are utilized to establish the timing of exposure. Both TLS and microtopographic profile gauges are used to obtain ground surface pro&#64257;les, from which Ex is estimated after the threshold distance (TD) is determined, i.e., the distance between the root and the sediment knickpoint, which allows de&#64257;ning the lowering of the ground surface caused by sheet erosion. For each profile, we measured the height between the topside of the root and a virtual plane tangential to the ground surface. In this way, we intended to avoid small-scale impacts of soil deformation, which may be due to pressures exerted by the root system, or by the arrangement of exposed roots. This may provoke small amounts of soil sedimentation or erosion depending on how they physically affect the surface runoff. We demonstrate that an adequate microtopographic characterization of exposed roots and their associated ground surface is very valuable to obtain accurate erosion rates. This finding could be utilized to develop the best management practices designed to eventually halt or perhaps, at least, lessen soil erosion, so that more sustainable management policies can be put into practice.

Characterizing erosion from dendrogeomorphology has usually focused on accurately finding the starting time of root exposure, by examining macroscopic or cell level changes caused by exposure. Here, we offer a detailed description of different novel techniques to obtain more precise erosion rates from highly accurate microtopographic data.

Sheet erosion is among the crucial drivers of soil degradation. Erosion is controlled by environmental factors and human activities, which often lead to ...

Measuring Stolons and Rhizomes of Turfgrasses Using a Digital Image Analysis System

Length and diameter of stolons or rhizomes are usually measured using simple rulers and calipers. This procedure is slow and laborious, so it is often used on a limited number of stolons or rhizomes. For this reason, these traits are limited in their use for morphological characterization of plants. The use of digital image analysis software technology may overcome measurement errors due to human mistakes, which tend to increase as the number and size of samples also increase. The protocol can be used for any kind of crop but is particularly suitable for forage or grasses, where plants are small and numerous. Turf samples consist of aboveground biomass and an upper soil layer to the depth of maximum rhizome development, depending on the species of interest. In studies, samples are washed from the soil, and stolons/rhizomes are cleaned by hand before analysis by digital image analysis software. The samples are further dried in a laboratory heating oven to measure dry weight; therefore, for each sample, the resultant data are total length, total dry weight, and average diameter. Scanned images can be corrected before analysis by excluding visible extraneous parts, such as remaining roots or leaves not removed with the cleaning process. Indeed, these fragments normally have much smaller diameters than stolons or rhizomes, so they can be easily excluded from analysis by fixing the minimum diameter below which objects are not considered. Stolon or rhizome density per unit area can then be calculated based on sample size. The advantage of this method is quick and efficient measurement of the length and average diameter of large sample numbers of stolons or rhizomes.

A software-based image analysis system provides an alternative method to study the morphology of stoloniferous and rhizomatous species. This protocol permits measurement of the length and diameter of stolons and rhizomes and can be applied to samples with a large amount of biomass and to a wide variety of species.

Length and diameter of stolons or rhizomes are usually measured using simple rulers and calipers. This procedure is slow and laborious, so it is often ...

Investigation of Plant Interactions Across Common Mycorrhizal Networks Using Rotated Cores

Arbuscular mycorrhizal (AM) fungi influence plant mineral nutrient uptake and growth, hence, they have the potential to influence plant interactions. The power of their influence is in extraradical mycelia that spread beyond nutrient depletion zones found near roots to ultimately interconnect individuals within a common mycorrhizal network (CMN). Most experiments, however, have investigated the role of AM fungi in plant interactions by growing plants with versus without mycorrhizal fungi, a method that fails to explicitly address the role of CMNs. Here, we propose a method that manipulates CMNs to investigate their role in plant interactions. Our method uses modified containers with conical bottoms with a nylon mesh and/or hydrophobic material covering slotted openings, 15N fertilizer, and a nutrient-poor interstitial sand. CMNs are left either intact between interacting individuals, severed by rotation of containers, or prevented from forming by a solid barrier. Our findings suggest that rotating containers is sufficient to disrupt CMNs and prevent their effects on plant interactions across CMNs. Our approach is advantageous because it mimics aspects of nature, such as seedlings tapping into already established CMNs and the use of a suite of AM fungi that may provide diverse benefits. Although our experiment is limited to investigating plants at the seedling stage, plant interactions across CMNs can be detected using our approach which therefore can be applied to investigate biological questions about the functioning of CMNs in ecosystems.

Most plants within communities likely are interconnected by arbuscular mycorrhizal (AM) fungi, but mediation of plant interactions by them has been investigated primarily by growing plants with versus without mycorrhizas. We present a method to manipulate common mycorrhizal networks among mycorrhizal plants to investigate their consequences for plant interactions.

Arbuscular mycorrhizal (AM) fungi influence plant mineral nutrient uptake and growth, hence, they have the potential to influence plant interactions. The ...

Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on 14NH4+/15NH4+ Analyses via Sequential Conversion to N2O

The importance of understanding the fate of nitrate (NO3&#8722;), which is the dominant N species transferred from terrestrial to aquatic ecosystems, has been increasing because global nitrogen loads have dramatically increased following industrialization. Dissimilatory nitrate reduction to ammonium (DNRA) and denitrification are both microbial processes that use NO3&#8722; for respiration. Compared to denitrification, quantitative determinations of the DNRA activity have been carried out only to a limited extent. This has led to an insufficient understanding of the importance of DNRA in NO3&#8722; transformations and the regulating factors of this process. The objective of this paper is to provide a detailed procedure for the measurement of the potential DNRA rate in environmental samples. In brief, the potential DNRA rate can be calculated from the 15N-labeled ammonium (15NH4+) accumulation rate in 15NO3&#8722; added incubation. The determination of the 14NH4+ and 15NH4+ concentrations described in this paper is comprised of the following steps. First, the NH4+ in the sample is extracted and trapped on an acidified glass filter as ammonium salt. Second, the trapped ammonium is eluted and oxidized to NO3&#8722; via persulfate oxidation. Third, the NO3&#8722; is converted to N2O via an N2O reductase deficient denitrifier. Finally, the converted N2O is analyzed using a previously developed quadrupole gas chromatography&#8211;mass spectrometry system. We applied this method to salt marsh sediments and calculated their potential DNRA rates, demonstrating that the proposed procedures allow a simple and more rapid determination compared to previously described methods.

Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on 14NH4+/15NH4+ Analyses via Sequential Conversion to N2O

A series of methods to determine the potential DNRA rate based on 14NH4+/15NH4+ analyses is provided in detail. NH4+ is converted into N2O via several steps and analyzed using quadrupole gas chromatography&#8211;mass spectrometry.

The importance of understanding the fate of nitrate (NO3&#8722;), which is the dominant N species transferred from terrestrial to aquatic ecosystems, has ...

Estimating Sediment Denitrification Rates Using Cores and N₂O Microsensors

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