Name: measurement of the potential rates of dissimilatory nitrate reduction to ammonium based on 14nh4+/15nh4+ analyses via sequential conversion to n2o
Uploaded: 2020-10-07T12:30:00.000+00:00
Duration: 8 min 5 s
Description: Watch this Scientific Journal Video about Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on 14NH4+/15NH4+ Analyses via Sequential Conversion to N2O at JoVE.com

We thank Naoto Tanaka for helping data collection and developing the protocol. The collection of samples was supported by JSPS KAKENHI Grant Number 17K15286.

1. Preparation of a PTFE envelope for quantitatively capturing gaseous NH3
<ol>
	<li>Place a 60-mm piece of polytetrafluoroethylene (PTFE) tape (25 mm in width) on a small sheet of aluminum foil (approximately 300 mm x 450 mm in size, wiped with ethanol).</li>
	<li>Ash a glass fiber filter (10 mm in diameter with a pore size of 2.7 &#956;m) at 450 &#176;C for 4 h in a muffle furnace. Place the glass fiber filter a little above the midpoint of the longer axis of the tape (Figure 1a).</li>
	<li>Spot 20 &#181;L of 0.9-mol/L H2SO4 on the center of a GF/D filter, and immediately fold the PTFE tape using two tweezers: flat-ended stamp tweezers and straight-ended tweezers. The following steps, steps 1.4&#8211;1.7, are shown in Figure 1 and should be conducted swiftly.</li>
	<li>Flip the PTFE tape over the GF/D filter at the dotted line shown in Figure 1a to form the shape shown in Figure 1b.</li>
	<li>Seal both sides by folding and then tightly pressing the edge with the tweezers (Figure 1c). Do not press too hard, and do not scratch the PTFE tape.</li>
	<li>Fold the open end with the tweezers, and then press the edge with the tweezers (Figure 1d).</li>
	<li>Seal the open end by tightly pressing the edge with the tweezers (Figure 1e). The GF/D filter should not be pressed during this procedure.</li>
</ol>
2. Preparing the biomass of a nitrous oxide reductase deficient denitrifier, Pseudomonas chlororaphis subsp. aureofaciens ATCC13985, for the denitrifier method
<ol>
	<li>Streak a 20% glycerol stock of Pseudomonas chlororaphis subsp. aureofaciens ATCC13985 on 1/4 strength tryptone soy broth (TSB) agar plates. Incubate the plates at 25 &#176;C for 2&#8211;3 days.</li>
	<li>Transfer a singles colony of P. chlororaphis to a small test tube containing 5 mL of autoclaved TSB medium and culture aerobically (without shaking) for a day at 25 &#176;C in dark until obtaining maximum growth; this will be used as the preculture.</li>
	<li>Transfer 3 mL of the preculture to a 1-L bottle with a silicon rubber stopper containing 1 L of freshly prepared autoclaved modified TSB supplemented with 10 mmol/L KNO323. Incubate the bottle while agitating using a stirrer under dark conditions at 25 &#176;C. After cultivating for 8 h, replace the silicon rubber stopper with a screw cap and close tightly. Continue the cultivation in anoxia overnight.</li>
	<li>Centrifuge the culture at 18,800 x g for 15 min at 4 &#176;C to obtain biomass pellets.</li>
	<li>Wash the packed biomass three times with 30 mL of Dulbecco&#8217;s phosphate-buffered saline (D-PBS(-), pH 7.5), to completely eliminate the NO3&#8722;. The conditions of the centrifugation are same as in Step 2.3.</li>
	<li>After washing, re-suspend the packed biomass with 30 mL of D-PBS(-). Use 1 mL of the suspension to determine the cell density by measuring OD600. Pipette 1 mL aliquots of the remaining suspension into sterile cryovials containing 0.8 mL of 45% glycerol. Preserve the prepared glycerol stocks at &#8722;80 &#176;C until use (see section 6).</li>
</ol>
3. Elimination of oxygen, nitrite, and nitrate from the sample sediment
<ol>
	<li>Weigh 3.0 g (wet weight) of sediment into a 20 mL glass vial and add 9.0 mL of surface water to suspend it (25% w/w slurry).</li>
	<li>Seal the vial with a black butyl rubber stopper (washed with ion-exchanged water and sterilized via autoclaving) and an aluminum cap.</li>
	<li>Purge the suspension and replace the headspace air with Ar (&#62;99.99%) at 0.6 L/min for 20 min using a manifold.</li>
	<li>Replace the Ar headspace gas with ultra-pure (&#62;99.99995%) He by vacuuming for 90 s and charging He for 30 s. Repeat this procedure four times. Set the headspace gas pressure to 1.5 atm.</li>
	<li>Incubate the vials at 20 &#176;C overnight with shaking at 150 rpm under dark conditions using a constant temperature shaker to eliminate the remaining oxygen, nitrate, and nitrite in the sediment suspension and headspace gas.</li>
</ol>
4. Time course experiment for determining DNRA rate
<ol>
	<li>Replace the headspace gas with fresh ultra-pure He using the same procedure as in step 3.5 but without the four repetitions.</li>
	<li>Add labeled and non-labeled substrates to each vial according to Table 1 through the butyl rubber stopper using a gastight syringe. Purge the previously prepared substrate solutions in adequately sized glass vials using ultra-pure He with the pressure of the headspace set to 1.5 atm to avoid air contamination. Avoid the unintentional injection of any amount of air during the injection procedures.</li>
	<li>Incubate vials at 20 &#176;C with shaking at 150 rpm. Add the substrates to each vial according to Table 1 and incubate for 1 h, 3 h, and 5 h. After incubation is halted, subject the sediment suspension in the vials to the following procedures: steps 4.4&#8211;4.9.</li>
	<li>Remove the aluminum cap and butyl rubber stopper from each vial. Then, add KCl (ashed at 450 &#176;C for 4 h) to the sediment suspension up to a final concentration of approximately 2 mol/L to ensure the recovery of NH4+ from the sediment. Close the vials with a butyl rubber stopper and an aluminum closure.</li>
	<li>Shake the sediment at 150 rpm for 1 h at 4 &#176;C under dark conditions to extract the NH4+.</li>
	<li>Transfer the entire sediment suspension in the vial to a 50 mL plastic centrifuge tube, and centrifuge at 10,000 x g for 10 min at 4 &#176;C.</li>
	<li>Rinse the internal wall of a freshly opened 10-mL disposable syringe, and attach it to a freshly opened disposable cellulose acetate membrane filter (pore size 0.22 &#181;m, 25 mm in diameter). Then, rinse the membrane filter with 1 mL of supernatant. Place the rinsed syringe&#8211;filter unit onto a 20-mL wide-mouth polypropylene (PP) bottle.</li>
	<li>Filter the remaining supernatant through an acetate membrane filter into the 20-mL PP bottle. Store the extracts extracts at &#8722;20 &#176;C until further analysis.</li>
</ol>
5. Capturing diffused NH4+ in 2M H2SO4 absorbed to the GF/D filter in the PTFE envelope and the persulfate oxidation of NH4+ to NO3&#8722;
<ol>
	<li>Prepare standard solutions of 14NH4Cl with concentration gradients of 0 &#956;mol/L, 10 &#956;mol/L, 40 &#956;mol/L, 100 &#956;mol/L, 200 &#956;mol/L, 400 &#956;mol/L, and 500 &#956;mol/L. For the 15N ratio analysis, fix the total concentration of NH4+ to 200 &#956;mol/L and prepare isotope ratios of 100:0, 99.5:0.5, 99:1, 93:7, 90:10, 50:50, and 10:90 with pure 14NH4Cl and 15NH4Cl standard solutions.</li>
	<li>Transfer 30 mg of MgO (ashed at 450 &#176;C for 4 h) to a 20-mL glass vial, and place the PTFE envelope in the vial.</li>
	<li>Transfer 5 mL of a sample or standard into the vial containing the MgO and the PTFE envelope, and immediately close with a gray butyl rubber stopper. Seal with an aluminum cap. If the concentration of NH4+ is expected to exceed 500 &#956;mol/L, dilute the sample to below 500 &#956;mol/L.</li>
	<li>Shake the vials at 150 rpm for 3 h at 4 &#176;C under dark conditions.</li>
	<li>Remove the aluminum cap and the butyl rubber stopper. Take the PTFE envelope out of the vial using point-ended tweezers, thoroughly rinse the envelope and the tweezers with ion-exchanged water, wipe them with a wiping paper, and then place the envelope on a fresh wiping paper.</li>
	<li>Open the PTFE envelope with a couple of tweezers (use of both the flat-ended and point-ended tweezers is recommended) in the exact reverse order of the folding performed in steps 1.4&#8211;1.7.</li>
	<li>Hold the peripheral area of the GF/D filter, where the H2SO4 is supposed to be unabsorbed, with the flat-ended tweezers, and transfer it into an 11-mL screw cap test tube with a PTFE -lined cap. Rinse the tweezers with ion-exchanged water, and wipe them with wiping paper.</li>
	<li>Repeat steps 5.5&#8211;5.7 for the remaining envelopes.</li>
	<li>Add 1 mL of ion-exchanged water to each of the test tubes, close the screw cap, and maintain it without shaking for at least 30 min at room temperature to completely elute the NH4+ from the GF/D filter. During this step, carry out the following step (step 5.10) in parallel.</li>
	<li>Prepare the persulfate-oxidizing solution (POR) reagent25,26.
	<ol>
		<li>Because it cannot be stored, prepare the exact amount of POR needed for treating a single day of samples.</li>
		<li>To prepare POR to treat 50 samples, pour 100 mL of ion-exchanged water into a 200-mL screw cap bottle and add 1.52 g of NaOH (nitrogen compound analysis grade), 3 g of boric acid, and 5 g of K2S2O8 (nitrogen and phosphorus analysis grade) in that order. Immediately after adding each reagent, shake the solution until it is completely dissolved.</li>
		<li>If necessary, soak the bottle in warm water to help the dissolution of the chemicals; however, attention should be paid with respect to the contamination of NO3&#8722; because tap water normally contains NO3&#8722;.</li>
	</ol>
	</li>
	<li>After Step 5.8, open the screw cap, add 2 mL of the POR reagent to the test tube, and close the tube tightly with a screw cap to prevent any loss or contamination during the following steps.</li>
	<li>Stand the test tubes on a rack, wrap them in double-layered aluminum foil, and autoclave them for 1 h at 121 &#176;C. Keep the tubes in an upright position during this step and avoid rapid changes in temperature after finishing the autoclaving.</li>
</ol>
6. Determining the NO3&#8722; converted from NH4+ by the denitrifier method using quadrupole GC/MS
<ol>
	<li>Mix 100 mL of sterile 40-mmol/L phosphate buffer (pH 7.2) and 100 mL of sterile 30-mmol/L glucose aseptically (20-mmol/L phosphate and 15-mmol/L glucose; final).</li>
	<li>Add a 1/7.2 volume of glycerol stock of P. chlororaphis to 200 mL of the phosphate-buffered glucose solution in a 300-mL Erlenmeyer flask, and purge with an ultra-pure He (&#62;99.995%) stream for 1 h.</li>
	<li>Dispense 2.0 mL of the denitrifier suspension into each 5-mL vial. Cap the vials with a gray butyl rubber stopper and an aluminum closure.</li>
	<li>Replace the headspace air with ultra-pure He by vacuuming for 3 min and charging the He for 1 min. Set the headspace gas positive pressure to 1.3 atm to avoid unintentional air contamination.</li>
	<li>Inject 1 mL of a sample or standard through the butyl rubber stopper using a 1-mL disposable syringe. Note the exact amount of the sample actually injected.</li>
	<li>Incubate the vials overnight at 25 &#176;C under dark conditions.</li>
	<li>Inject 0.3 mL of 6-mol/LL NaOH to stop the denitrification and absorb the headspace CO2, which will otherwise seriously disturb the N2O analysis by GC/MS because CO2 and N2O have the same molecular weight.</li>
	<li>Determine the amounts of 44N2O, 45N2O, and 46N2O in the headspace gas using quadrupole GC/MS with a modified injection port25. The operating conditions used for the GC/MS analysis are shown in Table 2.</li>
</ol>
7. Data analysis
<ol>
	<li>Derive the calibration curve for the NH4+ concentration from the linear relationship between the known concentration of 14NH4+ and the measured signal intensities of the total produced 44N2O + 45N2O + 46N2O. Derive the calibration curve for the 15N content from the linear relationship between the known atom% (i.e., 15N/14N+15N) and the calculated atom% using a previously provided equation27. Calculate the concentration of 15NH4+ by multiplying the total NH4+ by the 15N ratio of NH4+.</li>
	<li>Calculate the potential DNRA rate using equations provided elsewhere28,29,30.</li>
</ol>

The authors have nothing to disclose.

The concentration and isotope ratio of NH4+ for the DNRA analysis was quantified using several methods. The concentrations and isotope ratios of NH4+ are generally measured separately. The NH4+ concentration is typically measured using colorimetric methods including an autoanalyzer4,10,15,16,17. The isotope ratio measurement has wide variations depending on its method of NH4+ conversion, trapping, and instrumentation for the analysis. Typical methods include the following:
(1) The NH4+ in the sample is converted to NH3 via the addition of MgO or NaOH. After moving from the liquid phase to the gas phase, the NH3 is trapped on an acidified glass filter or in an acid solution. After drying, the filter is combusted and analyzed as N2 using an elemental analyzer/isotope ratio mass spectrometer (EA-IRMS)11,18,22,38. Alternatively, the captured NH4+ in an acid solution is collected on an adsorbent (e.g., zeolite) and is combusted and analyzed via EA-IRMS10,12.
(2) NH4+ is oxidized to N2 via hypobromite oxidation. The isotopic composition of the evolved N2 is measured via IRMS4,14,39 or membrane-introduction mass spectrometry (MIMS)16,17.
(3) The NH4+ concentration is determined using high-performance liquid chromatography without any conversion. Because the equilibrium of NH4+ and NH3 is slightly different for 15NH4+ and 14NH4+ near the pH of pKa, the 14NH4+ and 15NH4+ concentrations can be determined based on a small shift in the retention time6,19,40.
The method described in this paper is basically the same as approach (1) listed above, i.e., the step recovering NH4+ as NH3 on a glass filter under alkaline conditions; however, it is different with respect to the following sequential NH4+ conversion steps. These conversion steps are based on previous studies with added modifications to shorten the required time for a series of experiments. First, we made a few modifications to the original denitrifier method. After the biomass of P. chlororaphis is prepared as previously described23,24, we grow bacteria and preserve the concentrated cell suspension as a glycerol stock. This dense cell suspension can be directly used for the isotope analysis by mixing it with a buffer solution because the denitrifying activity has already been sufficiently induced. Even though further investigation is required, this modification may improve the reproducibility of the analysis because the presented method enables the mass-cultivation of denitrifier cells, which are directly available for analyses. We also modified the composition of the solution for suspending the bacterial cells, from a medium based on TSB to a phosphate-buffered glucose solution, to exclude the unnecessary components such as peptone in the original medium. This modification may reduce contamination by blank N because the phosphate-buffered glucose solution does not contain N, unlike the original medium used in previous studies; this should be tested via further analyses. In the step collecting NH4+ via the diffusion method, we shortened the incubation period and lowered the temperature to minimize the breakdown of organic N and any unfavorable conversion or loss of NH4+. The validity of this modification was checked using the linearity of the calibration curve. We also checked that the modified temperature and incubation period did not affect the recovery of NH4+ (data not shown).
Another advantage of this method is that NH4+ is ultimately converted to N2O, which has a low atmospheric background and can be measured using quadrupole GC/MS, which is less expensive and easier to manage than IRMS. Under the condition shown in table 2, CO2 (m/z 44) and N2O (m/z 44) are completely separated by GC; the retention time of these gases are 1.15 and 1.07 min, respectively. Since atmospheric concentration of N2O is on the order of ppb, N2O can be measured with negligible air interference even in low concentrations. The calibration curve of N2O passes almost through the origin, demonstrating the influence on the concentration of N2O due to atmospheric contamination is very limited (Figure 3). This method also has the advantage that it can quantify the 14NH4+ and 15NH4+ concentrations together; canonical methods, except approach (3) listed above, require individual analyses for the concentration and the isotope ratio.
Overall, the limits of detection and quantification for NH4+ using this method were approximately 0.03 &#956;mol and 0.09 &#956;mol, respectively, and these values are equivalent to 6 &#956;mol/L and 18 &#956;mol/L, if 5 mL of the sample solution (i.e., the sediment extract in this case) is used for the diffusion procedure as described in this paper. Even though using the colorimetric method is recommended to determine the NH4+ concentration of samples that have low NH4+, the proposed method effectively determines the NH4+ isotope ratio and the concentration in samples with relatively high concentrations of ammonium.

The artificial synthesis of nitrogen fertilizer and its widespread application have greatly perturbed the global nitrogen cycle. It is estimated that the transfer of reactive nitrogen from terrestrial to coastal systems has doubled since pre-industrial times1. A significant portion of fertilizers applied to a given field is washed away from the soil to rivers or groundwater, primarily as NO3&#8722; 2. This may cause environmental problems such as drinking water pollution, eutrophication, and the formation of hypoxia. NO3&#8722; in water environments is removed from or retained in the ecosystem via biological assimilation and various microbial dissimilatory processes. Denitrification and anammox are known to be major microbial removal processes for NO3&#8722;. Denitrification is the microbial reduction of NO3&#8722; to gaseous N products (NO, N2O, and N2) coupled with the oxidation of an electron donor, such as organic substances, thereby reducing the risk of the above-mentioned problems. Anammox also produces N2 from NO2&#8722; and NH4+; therefore, it removes inorganic N from an ecosystem. Conversely, DNRA works to retain N in an ecosystem; it is generally accepted that DNRA is performed primarily by fermentative bacteria or chemolithoautotrophic bacteria and that they reduce dissimilatory NO3&#8722; to bioavailable and less mobile NH4+.
Studies on DNRA have primarily been performed in marine or estuarine ecosystems, such as oceanic or estuarine sediments and water, salt or brackish marsh soil, and mangrove soil. Coastal or marine ecosystems are important as reservoirs for removing NO3&#8722; from terrestrial ecosystems, and in previous studies DNRA has been shown to contribute over a very wide range of NO3&#8722; removal (0&#8211;99%)3,4,5,6,7,8,9,10,11,12,13,14,15,16,17,18. Further, the existence of DNRA has been demonstrated in a wide range of environments including freshwater environments19, rice paddy soils20, and forest soils21. While these studies have shown that DNRA is potentially comparable to denitrification for NO3&#8722; removal, studies measuring the DNRA activity are still very limited compared to those measuring denitrification.
The DNRA rate has been evaluated using 15N-labeling techniques in conjunction with data analysis via analytical or numerical models. One analytical solution to calculate the DNRA rate is based on the increase in the 15N enrichment of the NH4+ pool after the addition of 15NO3&#8722; as a tracer. 15N-labeled NO3&#8722; is added to a sample and incubated, and the DNRA rate can then be calculated from the concentration and isotope ratio changes in NH4+ before and after a certain period of time. In this paper, a method to quantify the NH4+ concentration and the isotope ratio, which are required to calculate the DNRA rate, is described in detail. Basically, the method reported here is a combination of several previously reported techniques22,23,24,25,26 with modifications added to some procedures. The method is comprised of a series of five component procedures: (1) incubation of an environmental sample with the amendment of a stable isotope tracer, 15NO3&#8722;, (2) extraction and recovery of NH4+ using a &#8220;diffusion procedure&#8221; with modifications, (3) persulfate oxidation of NH4+ in the sample, consisting of indigenous NH4+ and 15NH4+ derived from 15NO3&#8722; via DNRA activity, into NO3&#8722; and 15NO3&#8722;, (4) subsequent microbial transformation of NO3&#8722; and 15NO3&#8722; to N2O isotopomers via the modified denitrifier method, and (5) quantification of the N2O isotopomers using gas chromatography&#8211;mass spectrometry (GC/MS). In the following section, first, the preparation for procedures (2) and (4) is described and then, subsequently, all five component procedures are described in detail.

<table><tbody><tr><td>&lt;sup&gt;15&lt;/sup&gt;N-KNO&lt;sub&gt;3&lt;/sub&gt;</td><td>SHOKO SCIENCE</td><td>N15-0197</td><td></td></tr><tr><td>&lt;sup&gt;15&lt;/sup&gt;N-NH&lt;sub&gt;4&lt;/sub&gt;Cl</td><td>SHOKO SCIENCE</td><td>N15-0034</td><td></td></tr><tr><td>20 mL PP bottle</td><td>SANPLATEC</td><td>61-3210-18</td><td>Wide-mouth</td></tr><tr><td>Aluminum cap</td><td>Maruemu</td><td>1307-13</td><td>No. 20, with hole</td></tr><tr><td>Boric acid</td><td>Wako</td><td>021-02195</td><td></td></tr><tr><td>Centrifuge</td><td>HITACHI</td><td>Himac CR21G II</td><td></td></tr><tr><td>Deoxygenized Gas Pressure &amp;amp; Replace Injector</td><td>SANSIN INDUSTRIAL</td><td>IP-12</td><td></td></tr><tr><td>Disposable cellulose acetate membrane filter</td><td>ADVANTEC</td><td>25CS020AS</td><td>Pore size 0.22 &amp;micro;m, 25 mm in diameter</td></tr><tr><td>Disposable syringe</td><td>Termo</td><td>SS-10SZ</td><td>10 mL</td></tr><tr><td>Disposable syringe</td><td>Termo</td><td>SS-01T</td><td>1 mL</td></tr><tr><td>Dulbecco&amp;rsquo;s Phosphate Buffered Saline (-)</td><td>NISSUI PHARMACEUTICAL</td><td>5913</td><td></td></tr><tr><td>Gastight syringe</td><td>VICI Valco Instruments</td><td>4075-15010</td><td>Series A-2, 100 &amp;micro;L</td></tr><tr><td>GC/MS</td><td>shimadzu</td><td>GCMS-QP2010ultra</td><td></td></tr><tr><td>GF/D</td><td>Whatman</td><td>1823-010</td><td>10 mm in diameter</td></tr><tr><td>Glass vial</td><td>Maruemu</td><td>0501-06</td><td>20 mL</td></tr><tr><td>Gray butyl rubber stopper</td><td>Maruemu</td><td>1306-03</td><td>No.20-S</td></tr><tr><td>H&lt;sub&gt;2&lt;/sub&gt;SO&lt;sub&gt;4&lt;/sub&gt;</td><td>Wako</td><td>192-04696</td><td>Guaranteed Reagent</td></tr><tr><td>K&lt;sub&gt;2&lt;/sub&gt;S&lt;sub&gt;2&lt;/sub&gt;O&lt;sub&gt;8&lt;/sub&gt;</td><td>Wako</td><td>169-11891</td><td>Nitrogen and Phosphorus analysis grade</td></tr><tr><td>KCl</td><td>Wako</td><td>163-03545</td><td>Guaranteed Reagent</td></tr><tr><td>KNO&lt;sub&gt;3&lt;/sub&gt;</td><td>Wako</td><td>160-04035</td><td>Guaranteed Reagent</td></tr><tr><td>NaOH</td><td>Wako</td><td>191-08625</td><td>Nitrogen compounds analysis grade</td></tr><tr><td>NH&lt;sub&gt;4&lt;/sub&gt;Cl</td><td>Wako</td><td>017-02995</td><td>Guaranteed Reagent</td></tr><tr><td>Plastic centrifuge tube</td><td>ASONE</td><td>1-3500-22</td><td>50 mL, VIO-50BN</td></tr><tr><td>Pseudomonas chlororaphis subsp. aureofaciens</td><td>American Type Culture Collection (ATCC)</td><td>ATCC 13985</td><td>Freeze-dried, the type strain of Pseudomonas aureofaciens</td></tr><tr><td>PTFE sealing tape</td><td>Sigma-Aldrich</td><td>Z221880</td><td>25 mm in width</td></tr><tr><td>Reciprocating shaker</td><td>TAITEC</td><td>0000207-000</td><td>NR-10</td></tr><tr><td>Screw-cap test tube</td><td>IWAKI</td><td>84-0252</td><td>11 mL</td></tr><tr><td>PTFE-lined cap for test tube</td><td>IWAKI</td><td>84-0262</td><td></td></tr><tr><td>Tryptic Soy Broth</td><td>Difco Laboratories</td><td>211825</td><td></td></tr></tbody></table>

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.

The representative results presented in this paper were derived from 15N-tracing experiments of salt marsh sediments. The sampled salt marsh was newly created in the aftermath of the 2011 Great East Japan Earthquake in the Moune area of Kesen-numa city in Miyagi Prefecture, Japan. In September 2017, surface sediments (0&#8211;3 cm) were collected at two sites in the subtidal and intertidal zones. First, immediately after collection, the sediment was sieved through a 4-mm mesh to remove plant roots, shells debris, and rubble and then homogenized. The samples were stored at 4 &#176;C until the DNRA analysis was conducted.
The incubation procedures for the 15NO3&#8722; and the simultaneous determination of the 14NH4+ and 15NH4+ concentrations were carried out as described in the protocol section. An increase in the 15NH4+ concentration throughout the incubation period was observed for all sediments (Figure 2). We calculated the DNRA rates by dividing the accumulation rate of 15NH4+ by the isotope ratio of the NO3&#8722; pool29. The calculated rates were within the range of 24.8&#8211;177 nmol-N g&#8722;1 dry soil h&#8722;1 (Table 3) and were comparable to values found in previous studies. This range of obtained rates is higher than the reported values derived from similar environments including those from intertidal sediments17, salt marshes5,16, and other estuarine environments18,33,34, as well as from eutrophic environments such as a shallow river estuary in North Carolina31 and the Shanghai urban river networks32. Conversely, Fernandes et al.13 reported higher potential DNRA rates in organically rich mangrove soils in India. In general, DNRA is thought to be favored by a high ratio of available C to electron acceptors35,36,37. The samples demonstrating the representative results were taken from a salt marsh newly created by an earthquake, which had originally been used as a cultivation field. This particular characteristic of the samples may contribute to the observed high DNRA rate. Consistent with this speculation, the DNRA rate in the intertidal zone, which is rich in organic compounds (data not shown) compared to the subtidal zone, was higher than that in the subtidal zone (Figure 2, Table 3).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59562/59562fig1.jpg" /> 
Figure 1: Preparation of a PTFE envelope for capturing gaseous NH3. The PTFE envelope used in the diffusion procedure is prepared by folding PTFE tape following the instructions shown in panels (A)&#8211;(E). The acidified filter inside the envelope captures the gaseous NH3. These steps should be conducted quickly. Detailed information is given in steps 1.2&#8211;1.7 in the protocol section. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59562/59562fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/59562/59562fig2.jpg" /> 
Figure 2: Change in the 15NH4+ concentration via the anaerobic incubation of sediments. The 15N tracer incubations of the sediment samples were conducted in duplicate. The concentration of 15N-NH4+ is shown in nmol per dry weight of sediment. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59562/59562fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/59562/59562fig3.jpg" /> 
Figure 3: An example of calibration curve of low concentration N2O. Peak area of N2O was obtained by sum of the peak area of m/z 44, m/z 45, and m/z 46. Configurations for GC/MS analysis is shown in Table 2. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59562/59562fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td colspan="2">15N-labeled and non-labeled substrates added to each vial</td>
		</tr>
		<tr>
			<td></td>
			<td>100mM</td>
			<td>100mM</td>
		</tr>
		<tr>
			<td></td>
			<td>NH4Cl</td>
			<td>K15NO3</td>
		</tr>
		<tr>
			<td>Volume (&#181;L) of 100 mM stock solution added to each vial</td>
			<td>24</td>
			<td>60</td>
		</tr>
		<tr>
			<td>Final concentration (&#181;mol/L)*</td>
			<td>&#62;230&#167;</td>
			<td>570</td>
		</tr>
		<tr>
			<td colspan="3">*shown values are calculated by assuming that water content of the sediment is 50%&#160;</td>
		</tr>
		<tr>
			<td colspan="3">&#167;depending on the background ammonium concentration</td>
		</tr>
	</tbody>
</table>
Table 1: Combinations of substrates amended to vials retaining approximately 11 mL of the sediment suspension. Samples were prepared in duplicate and subjected to further analyses after 1 h, 3 h, and 5 h of incubation.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Equipment</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;&#160; quadrupole GCMS</td>
			<td>Shimadzu GCMS-QP2010 Ultra</td>
		</tr>
		<tr>
			<td>&#160;&#160; column</td>
			<td>CP-PoraBONDQ 25m; &#966; 0.32mm; film thickness, 5&#181;m</td>
		</tr>
		<tr>
			<td>Analytical conditions</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160; column temp</td>
			<td>40 &#176;C</td>
		</tr>
		<tr>
			<td>&#160; injection port temp</td>
			<td>100 &#176;C</td>
		</tr>
		<tr>
			<td>&#160; carrier gas stream</td>
			<td>Total flow rate, 47.1 mL&#8226;min-1 
			flow rate in column, 2.10 mL&#8226;min-1</td>
		</tr>
		<tr>
			<td>&#160; sprit ratio</td>
			<td>20</td>
		</tr>
		<tr>
			<td>&#160; detection voltage</td>
			<td>1.5 kv</td>
		</tr>
		<tr>
			<td>Sensitivity of N2O</td>
			<td></td>
		</tr>
		<tr>
			<td>&#160;&#160; lower limit of detection (LOD)*</td>
			<td>1.42 pmol</td>
		</tr>
		<tr>
			<td>&#160;&#160; lower limit of quantification (LOQ)*</td>
			<td>4.58 pmol</td>
		</tr>
		<tr>
			<td>*LOD and LOQ were determined by a linear relationship among a serial dilution of N2O (0.97, 1.94, 2.91, 4.75, 9.50, 14.3 ppm) in He, corresponding responses in peak area, and S/N ratio. LOD and LOQ were calculated as concentrations equivalent to S/N=3 and S/N=10, respectively.</td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 2: Conditions for the GC/MS analysis.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>sediment</td>
			<td>DNRA rate</td>
			<td>&#160;enrichment of NO3- *</td>
		</tr>
		<tr>
			<td></td>
			<td>nmol-N g-1 hr-1</td>
			<td>atom%</td>
		</tr>
		<tr>
			<td>Intertidal 1</td>
			<td>177</td>
			<td>99.9</td>
		</tr>
		<tr>
			<td>Intertidal 2</td>
			<td>129</td>
			<td>99.0</td>
		</tr>
		<tr>
			<td>Subtidal 1</td>
			<td>39.3</td>
			<td>99.9</td>
		</tr>
		<tr>
			<td>Subtidal 2</td>
			<td>24.8</td>
			<td>99.0</td>
		</tr>
		<tr>
			<td>*same as atom% of added KNO3; complete elimination and no nitrification under used incubation conditions was tested&#160; previously.&#160;</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 3: Potential DNRA rates of the tested intertidal and subtidal sediments.

Watch this Scientific Journal Video about Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on 14NH4+/15NH4+ Analyses via Sequential Conversion to N2O at JoVE.com

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.

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 ...

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Nanyang Technological University

Research

JoVE Journal

Environment

6.0K Views.  Chuo University. Kust- en mariene ecosystemen zijn belangrijk als reservoirs, voor het verwijderen van nitraat uit terrestrische ecosystemen. Nitraat in het aquatisch milieu kan door meerdere processen tegelijk worden geconsumeerd. Zoals denitrification, anammox en DNRA.Hoewel eerdere studies hebben aangetoond dat DNRA mogelijk schuldig is aan denitrificatie, studies meten DNRA activiteit als nog steeds zeer beperkt in vergelijking met die meten denitrificatie. In ons protocol bieden we een gedetaillee...

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

A General Method for Detecting Nitrosamide Formation in the In Vitro Metabolism of Nitrosamines by Cytochrome P450s

N-nitrosamines are a well-established group of environmental carcinogens, which require cytochrome P450 oxidation to exhibit activity. The accepted mechanism of metabolic activation involves formation of &#945;-hydroxynitrosamines that spontaneously decompose to DNA alkylating agents. Accumulation of DNA damage and the resulting mutations can ultimately lead to cancer. New evidence indicates that &#945;-hydroxynitrosamines can be further oxidized to nitrosamides processively by cytochrome P450s. Because nitrosamides are generally more stable than &#945;-hydroxynitrosamines and can also alkylate DNA, nitrosamides may play a role in carcinogenesis. In this report, we describe a general protocol for evaluating nitrosamide production from in vitro cytochrome P450-catalyzed metabolism of nitrosamines. This protocol utilizes a general approach to the synthesis of the relevant nitrosamides and an in vitro cytochrome P450 metabolism assay using liquid chromatography-nanospray ionization-high resolution tandem mass spectrometry for detection. This method detected N&#8242;-nitrosonorcotinine as a minor metabolite of N&#8242;-nitrosonornicotine in the example study. The method has high sensitivity and selectively due to accurate mass detection. Application of this method to a wide variety of nitrosamine-cytochrome P450 systems will help determine the generality of this transformation. Because cytochrome P450s are polymorphic and vary in activity, a better understanding of nitrosamide formation could aid in individual cancer risk assessment.

A General Method for Detecting Nitrosamide Formation in the In Vitro Metabolism of Nitrosamines by Cytochrome P450s

&#945;-hydroxylation of carcinogenic nitrosamines by cytochrome P450s is the accepted metabolic pathway that produces DNA-damaging intermediates, which cause mutations. However, new data indicates further oxidation to nitrosamides can occur. We describe a general method for detecting nitrosamides produced from in vitro cytochrome P450-catalyzed metabolism of nitrosamines.

N-nitrosamines are a well-established group of environmental carcinogens, which require cytochrome P450 oxidation to exhibit activity. The accepted ...

Cancer Research

Calibrated Passive Sampling - Multi-plot Field Measurements of NH3 Emissions with a Combination of Dynamic Tube Method and Passive Samplers

Agricultural ammonia (NH3) emissions (90% of total EU emissions) are responsible for about 45% airborne eutrophication, 31% soil acidification and 12% fine dust formation within the EU15. But NH3 emissions also mean a considerable loss of nutrients. Many studies on NH3 emission from organic and mineral fertilizer application have been performed in recent decades. Nevertheless, research related to NH3 emissions after application fertilizers is still limited in particular with respect to relationships to emissions, fertilizer type, site conditions and crop growth. Due to the variable response of crops to treatments, effects can only be validated in experimental designs including field replication for statistical testing. The dominating ammonia loss methods yielding quantitative emissions require large field areas, expensive equipment or current supply, which restricts their application in replicated field trials. This protocol describes a new methodology for the measurement of NH3 emissions on many plots linking a simple semi-quantitative measuring method used in all plots, with a quantitative method by simultaneous measurements using both methods on selected plots. As a semi-quantitative measurement method passive samplers are used. The second method is a dynamic chamber method (Dynamic Tube Method) to obtain a transfer quotient, which converts the semi-quantitative losses of the passive sampler to quantitative losses (kg nitrogen ha-1). The principle underlying this approach is that passive samplers placed in a homogeneous experimental field have the same NH3 absorption behavior under identical environmental conditions. Therefore, a transfer co-efficient obtained from single passive samplers can be used to scale the values of all passive samplers used in the same field trial. The method proved valid under a wide range of experimental conditions and is recommended to be used under conditions with bare soil or small canopies (&#60;0.3 m). Results obtained from experiments with taller plants should be treated more carefully.

Calibrated Passive Sampling - Multi-plot Field Measurements of NH3 Emissions with a Combination of Dynamic Tube Method and Passive Samplers

Ammonia emissions are a major threat to the environment by eutrophication, soil acidification and fine particle formation and stem mainly from agricultural sources. This method allows ammonia loss measurements in replicated field trials enabling statistical analysis of emissions and of relationships between crop development and emissions.

Agricultural ammonia (NH3) emissions (90% of total EU emissions) are responsible for about 45% airborne eutrophication, 31% soil acidification and 12% ...

Electrochemically and Bioelectrochemically Induced Ammonium Recovery

Streams such as urine and manure can contain high levels of ammonium, which could be recovered for reuse in agriculture or chemistry. The extraction of ammonium from an ammonium-rich stream is demonstrated using an electrochemical and a bioelectrochemical system. Both systems are controlled by a potentiostat to either fix the current (for the electrochemical cell) or fix the potential of the working electrode (for the bioelectrochemical cell). In the bioelectrochemical cell, electroactive bacteria catalyze the anodic reaction, whereas in the electrochemical cell the potentiostat applies a higher voltage to produce a current. The current and consequent restoration of the charge balance across the cell allow the transport of cations, such as ammonium, across a cation exchange membrane from the anolyte to the catholyte. The high pH of the catholyte leads to formation of ammonia, which can be stripped from the medium and captured in an acid solution, thus enabling the recovery of a valuable nutrient. The flux of ammonium across the membrane is characterized at different anolyte ammonium concentrations and currents for both the abiotic and biotic reactor systems. Both systems are compared based on current and removal efficiencies for ammonium, as well as the energy input required to drive ammonium transfer across the cation exchange membrane. Finally, a comparative analysis considering key aspects such as reliability, electrode cost, and rate is made.
This video article and protocol provide the necessary information to conduct electrochemical and bioelectrochemical ammonia recovery experiments. The reactor setup for the two cases is explained, as well as the reactor operation. We elaborate on data analysis for both reactor types and on the advantages and disadvantages of bioelectrochemical and electrochemical systems.

We demonstrate the extraction of ammonium from an ammonium-rich stream using an electrochemical and a bioelectrochemical system. The reactor setup, operation and data analysis are discussed.

Streams such as urine and manure can contain high levels of ammonium, which could be recovered for reuse in agriculture or chemistry. The extraction of ...

Chemistry

The Benthic Exchange of O2, N2 and Dissolved Nutrients Using Small Core Incubations

The measurement of sediment-water exchange of gases and solutes in aquatic sediments provides data valuable for understanding the role of sediments in nutrient and gas cycles. After cores with intact sediment-water interfaces are collected, they are submerged in incubation tanks and kept under aerobic conditions at in situ temperatures. To initiate a time course of overlying water chemistry, cores are sealed without bubbles using a top cap with a suspended stirrer. Time courses of 4-7 sample points are used to determine the rate of sediment water exchange. Artificial illumination simulates day-time conditions for shallow photosynthetic sediments, and in conjunction with dark incubations can provide net exchanges on a daily basis. The net measurement of N2 is made possible by sampling a time course of dissolved gas concentrations, with high precision mass spectrometric analysis of N2:Ar ratios providing a means to measure N2 concentrations. We have successfully applied this approach to lakes, reservoirs, estuaries, wetlands and storm water ponds, and with care, this approach provides valuable information on biogeochemical balances in aquatic ecosystems.

The Benthic Exchange of O2, N2 and Dissolved Nutrients Using Small Core Incubations

Here, we present small core incubations for the measurement of sediment-water gas and solute exchange. These will provide reliable measurements of sediment-water exchange that assess the role of sediment in influencing biological and biogeochemical processes in aquatic ecosystems.

The measurement of sediment-water exchange of gases and solutes in aquatic sediments provides data valuable for understanding the role of sediments in ...

Automated, High-resolution Mobile Collection System for the Nitrogen Isotopic Analysis of NOx

Nitrogen oxides (NOx = NO + NO2) are a family of atmospheric trace gases that have great impact on the environment. NOx concentrations directly influence the oxidizing capacity of the atmosphere through interactions with ozone and hydroxyl radicals. The main sink of NOx is the formation and deposition of nitric acid, a component of acid rain and a bioavailable nutrient. NOx is emitted from a mixture of natural and anthropogenic sources, which vary in space and time. The collocation of multiple sources and the short lifetime of NOx make it challenging to quantitatively constrain the influence of different emission sources and their impacts on the environment. Nitrogen isotopes of NOx have been suggested to vary amongst different sources, representing a potentially powerful tool to understand the sources and transport of NOx. However, previous methods of collecting atmospheric NOx integrate over long (week to month) time spans and are not validated for the efficient collection of NOx in relevant, diverse field conditions. We report on a new, highly efficient field-based system that collects atmospheric NOx for isotope analysis at a time resolution between 30 min and 2 hr. This method collects gaseous NOx in solution as nitrate with 100% efficiency under a variety of conditions. Protocols are presented for collecting air in urban settings under both stationary and mobile conditions. We detail the advantages and limitations of the method and demonstrate its application in the field. Data from several deployments are shown to 1) evaluate field-based collection efficiency by comparisons with in situ NOx concentration measurements, 2) test the stability of stored solutions before processing, 3) quantify in situ reproducibility in a variety of urban settings, and 4) demonstrate the range of N isotopes of NOx detected in ambient urban air and on heavily traveled roadways.

Automated, High-resolution Mobile Collection System for the Nitrogen Isotopic Analysis of NOx

Previous work suggests that the nitrogen isotopic composition of atmospheric nitrogen oxides might distinguish the influence of different sources in the environment. We report on an automated, mobile, field-based method for the high collection efficiency of atmospheric NOx for N isotopic analysis at an hourly time resolution.

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Procedures of Laboratory Fumigation for Pest Control with Nitric Oxide Gas

Nitric oxide (NO) is a newly discovered fumigant for postharvest pest control. This paper provides detailed protocols for conducting NO fumigation on fresh products and procedures for residue analysis and product quality evaluation. An airtight fumigation chamber containing fresh fruit and vegetables is first flushed with nitrogen (N2) to establish an ultralow oxygen (ULO) environment followed by injection of NO. The fumigation chamber is then kept at a low temperature of 2 - 5 &#176;C for a specified time period necessary to kill a target pest to complete a fumigation treatment. At the end of a fumigation treatment, the fumigation chamber is flushed with N2 to dilute NO prior to opening the chamber to ambient air to prevent the reaction between NO and O2, which produces NO2 and may damage delicate fresh products. At different times after NO fumigation, NO2 in headspace and nitrate and nitrite in liquid samples were measured as residues. Product quality was evaluated after 2 weeks of post-treatment cold storage to determine effects of NO fumigation on product quality. Keeping&#160;O2 from reacting with NO is critical to NO fumigation and is an important part of the protocols. Measuring NO levels is challenging and a practical solution is provided. Possible protocol modifications are also suggested for measuring NO levels in the fumigation chambers as well as residues. NO fumigation has the potential to be a practical alternative to methyl bromide fumigation for postharvest pest control on fresh and stored products. This publication is intended to assist other researchers in conducting NO fumigation research for postharvest pest control and accelerating the development of NO fumigation for practical applications.

This paper describes nitric oxide (NO) fumigation protocols for postharvest pest control. Fumigation chambers are flushed with nitrogen (N2) to establish ultralow oxygen conditions before NO is injected. At the end, chambers are flushed with N2 to dilute NO before exposing products to ambient air to prevent exposure to NO2.

Nitric oxide (NO) is a newly discovered fumigant for postharvest pest control. This paper provides detailed protocols for conducting NO fumigation on ...

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.

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.

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

Microplot Design and Plant and Soil Sample Preparation for 15Nitrogen Analysis

Many&#160;nitrogen fertilizer studies evaluate the overall effect of a treatment on end-of-season measurements such as grain yield or cumulative N losses. A stable isotope approach is necessary to follow and quantify the fate of fertilizer derived N (FDN) through the soil-crop system. The purpose of this paper is to describe a small-plot research&#160;design utilizing non-confined 15N enriched microplots for multiple soil and plant sampling events over two growing seasons and provide sample collection, handling, and processing protocols for total 15N analysis. The methods were demonstrated using a replicated study from south-central Minnesota planted to corn (Zea mays L.). Each treatment consisted of six corn rows (76 cm row-spacing) 15.2 m long with a microplot (2.4 m by 3.8 m) embedded at one end. Fertilizer-grade urea was applied at 135 kg N&#8729;ha-1 at planting, while the microplot received urea enriched to 5 atom % 15N. Soil and plant samples were taken several times throughout the growing season, taking care to minimize cross-contamination by using separate tools and physically separating unenriched and enriched samples during all procedures. Soil and plant samples were dried, ground to pass through a 2 mm screen, and then ground to a flour-like consistency using a roller jar mill. Tracer studies require additional planning, sample processing time and manual labor, and incur higher costs for 15N enriched materials and sample analysis than traditional N studies. However, using the mass balance approach, tracer studies with multiple in-season sampling events allow the researcher to estimate FDN distribution through the soil-crop system and estimate unaccounted-for FDN from the system.

Microplot Design and Plant and Soil Sample Preparation for 15Nitrogen Analysis

A microplot design for 15N tracer research is described to accommodate multiple in-season plant and soil sampling events. Soil and plant sample collection and processing procedures, including grinding and weighing protocols, for 15N analysis are put forth.

Many&#160;nitrogen fertilizer studies evaluate the overall effect of a treatment on end-of-season measurements such as grain yield or cumulative N losses. ...

Workflow Based on the Combination of Isotopic Tracer Experiments to Investigate Microbial Metabolism of Multiple Nutrient Sources

Studies in the field of microbiology rely on the implementation of a wide range of methodologies. In particular, the development of appropriate methods substantially contributes to providing extensive knowledge of the metabolism of microorganisms growing in chemically defined media containing unique nitrogen and carbon sources. In contrast, the management through metabolism of multiple nutrient sources, despite their broad presence in natural or industrial environments, remains virtually unexplored. This situation is mainly due to the lack of suitable methodologies, which hinders investigations.
We report an experimental strategy to quantitatively and comprehensively explore how metabolism operates when a nutrient is provided as a mixture of different molecules, i.e., a complex resource. Here, we describe its application for assessing the partitioning of multiple nitrogen sources through the yeast metabolic network. The workflow combines information obtained during stable isotope tracer experiments using selected 13C- or 15N-labeled substrates. It first consists of parallel and reproducible fermentations in the same medium, which includes a mixture of N-containing molecules; however,a selected nitrogen source is labeled each time. A combination of analytical procedures (HPLC, GC-MS) is implemented to assess the labeling patterns of targeted compounds and to quantify the consumption and recovery of substrates in other metabolites. An integrated analysis of the complete dataset provides an overview of the fate of consumed substrates within cells. This approach requires an accurate protocol for the collection of samples&#8211;facilitated by a robot-assisted system for online monitoring of fermentations&#8211;and the achievement of numerous time-consuming analyses. Despite these constraints, it allowed understanding, for the first time, the partitioning of multiple nitrogen sources throughout the yeast metabolic network. We elucidated the redistribution of nitrogen from more abundant sources toward other N-compounds and determined the metabolic origins of volatile molecules and proteinogenic amino acids.

This protocol describes an experimental procedure to quantitatively and comprehensively investigate the metabolism of multiple nutrient sources. This workflow, based on a combination of isotopic tracer experiments and an analytical procedure, allows the fate of consumed nutrients and the metabolic origin of molecules synthetized by microorganisms to be determined.

Studies in the field of microbiology rely on the implementation of a wide range of methodologies. In particular, the development of appropriate methods ...

Bioengineering

Chemiluminescence-based Assays for Detection of Nitric Oxide and its Derivatives from Autoxidation and Nitrosated Compounds

Nitric oxide (NO) activity in vivo is the combined results of its direct effects, the action of its derivatives generated from NO autoxidation, and the effects of nitrosated compounds. Measuring NO metabolites is essential to studying NO activity both at vascular levels and in other tissues, especially in the experimental settings where exogenous NO is administered. Ozone-based chemiluminescence assays allow precise measurements of NO and NO metabolites in both fluids (including plasma, tissue homogenates, cell cultures) and gas mixtures (e.g., exhaled breath). NO reacts with ozone to generate nitrogen dioxide in an excited state. The consequent light emission allows photodetection and the generation of an electric signal reflecting the NO content of the sample. Aliquots from the same sample can be used to measure specific NO metabolites, such as nitrate, nitrite, S-nitrosothiols, and iron-nitrosyl complexes. In addition, NO consumed by cell-free hemoglobin is also quantified with chemiluminescence analysis. An illustration of all these techniques is provided.

Here, we present protocols for detecting nitric oxide and its biologically relevant derivatives using chemiluminescence-based assays with high sensitivity.

Nitric oxide (NO) activity in vivo is the combined results of its direct effects, the action of its derivatives generated from NO autoxidation, and the ...

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

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