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 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>15N-KNO3</td>
			<td>SHOKO SCIENCE</td>
			<td>N15-0197</td>
			<td></td>
		</tr>
		<tr>
			<td>15N-NH4Cl</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 &#38; 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 &#181;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&#8217;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 &#181;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>H2SO4</td>
			<td>Wako</td>
			<td>192-04696</td>
			<td>Guaranteed Reagent</td>
		</tr>
		<tr>
			<td>K2S2O8</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>KNO3</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>NH4Cl</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 t...

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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6.0K Views.  Chuo University. 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–mass spectrometry.

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

High-throughput Fluorometric Measurement of Potential Soil Extracellular Enzyme Activities

Microbes in soils and other environments produce extracellular enzymes to depolymerize and hydrolyze organic macromolecules so that they can be assimilated for energy and nutrients. Measuring soil microbial enzyme activity is crucial in understanding soil ecosystem functional dynamics. The general concept of the fluorescence enzyme assay is that synthetic C-, N-, or P-rich substrates bound with a fluorescent dye are added to soil samples. When intact, the labeled substrates do not fluoresce. Enzyme activity is measured as the increase in fluorescence as the fluorescent dyes are cleaved from their substrates, which allows them to fluoresce. Enzyme measurements can be expressed in units of molarity or activity. To perform this assay, soil slurries are prepared by combining soil with a pH buffer. The pH buffer (typically a 50 mM sodium acetate or 50 mM Tris buffer), is chosen for the buffer&#39;s particular acid dissociation constant (pKa) to best match the soil sample pH. The soil slurries are inoculated with a nonlimiting amount of fluorescently labeled (i.e. C-, N-, or P-rich) substrate. Using soil slurries in the assay serves to minimize limitations on enzyme and substrate diffusion. Therefore, this assay controls for differences in substrate limitation, diffusion rates, and soil pH conditions; thus detecting potential enzyme activity rates as a function of the difference in enzyme concentrations (per sample).
Fluorescence enzyme assays are typically more sensitive than spectrophotometric (i.e. colorimetric) assays,&#160;but can suffer from interference caused by impurities and the instability of many fluorescent compounds when exposed to light; so caution is required when handling fluorescent substrates. Likewise, this method only assesses potential enzyme activities under laboratory conditions when substrates are not limiting. Caution should be used when interpreting the data representing cross-site comparisons with differing temperatures or soil types, as in situ soil type and temperature can influence enzyme kinetics.

To measure potential rates of soil extracellular enzyme activities, synthetic substrates that are bound to a fluorescent dye are added to soil samples. Enzyme activity is measured as the fluorescent dye is released from the substrate by an enzyme-catalyzed reaction, where higher fluorescence indicates more substrate degradation.

Microbes in soils and other environments produce extracellular enzymes to depolymerize and hydrolyze organic macromolecules so that they can be ...

Methods for Facilitating Microbial Growth on Pulp Mill Waste Streams and Characterization of the Biodegradation Potential of Cultured Microbes

The kraft process is applied to wood chips for separation of lignin from the polysaccharides within lignocellulose for pulp that will produce a high quality paper. Black liquor is a pulping waste generated by the kraft process that has potential for downstream bioconversion. However, the recalcitrant nature of the lignocellulose resources, its chemical derivatives that constitute the majority of available organic carbon within black liquor, and its basic pH present challenges to microbial biodegradation of this waste material. Methods for the collection and modification of black liquor for microbial growth are aimed at utilization of this pulp waste to convert the lignin, organic acids, and polysaccharide degradation byproducts into valuable chemicals. The lignocellulose extraction techniques presented provide a reproducible method for preparation of lignocellulose growth substrates for understanding metabolic capacities of cultured microorganisms. Use of gas chromatography-mass spectrometry enables the identification and quantification of the fermentation products resulting from the growth of microorganisms on pulping waste. These methods when used together can facilitate the determination of the metabolic activity of microorganisms with potential to produce fermentation products that would provide greater value to the pulping system and reduce effluent waste, thereby increasing potential paper milling profits and offering additional uses for black liquor.

Industrial wastes can be collected and modified to analyze microbial growth. Lignocellulose extraction techniques provide components to analyze specific biodegradation ability. Gas chromatography-mass spectrometry identifies fermentation products of microorganisms grown on pulping waste. These methods determine the metabolic capacity of microorganisms to degrade pulping waste.

The kraft process is applied to wood chips for separation of lignin from the polysaccharides within lignocellulose for pulp that will produce a high ...

Measuring Carbon-based Contaminant Mineralization Using Combined CO2 Flux and Radiocarbon Analyses

A method is described which uses the absence of radiocarbon in industrial chemicals and fuels made from petroleum feedstocks which frequently contaminate the environment. This radiocarbon signal &#8212; or rather the absence of signal &#8212; is evenly distributed throughout a contaminant source pool (unlike an added tracer) and is not impacted by biological, chemical or physical processes (e.g., the 14C radioactive decay rate is immutable). If the fossil-derived contaminant is fully degraded to CO2, a harmless end-product, that CO2 will contain no radiocarbon. CO2 derived from natural organic matter (NOM) degradation will reflect the NOM radiocarbon content (usually &#60;30,000 years old). Given a known radiocarbon content for NOM (a site background), a two end-member mixing model can be used to determine the CO2 derived from a fossil source in a given soil gas or groundwater sample. Coupling the percent CO2 derived from the contaminant with the CO2 respiration rate provides an estimate for the total amount of contaminant degraded per unit time. Finally, determining a zone of influence (ZOI) representing the volume from which site CO2 is collected allows determining the contaminant degradation per unit time and volume. Along with estimates for total contaminant mass, this can ultimately be used to calculate time-to-remediate or otherwise used by site managers for decision-making.

Measuring Carbon-based Contaminant Mineralization Using Combined CO2 Flux and Radiocarbon Analyses

A protocol is described wherein CO2 mineralized from organic contaminant (derived from petroleum feedstocks) biodegradation is trapped, quantified, and analyzed for 14C content. A model is developed to determine CO2 capture zone's spatial extent. Spatial and temporal measurements allow integrating contaminant mineralization rates for predicting remediation extent and time.

A method is described which uses the absence of radiocarbon in industrial chemicals and fuels made from petroleum feedstocks which frequently contaminate ...

Assessment of Labile Organic Carbon in Soil Using Sequential Fumigation Incubation Procedures

Management practices and environmental changes can alter soil nutrient and carbon cycling. Soil labile organic carbon, a readily decomposable C pool, is highly sensitive to disturbance. It is also the primary substrate for soil microorganisms, which is fundamental to nutrient cycling. Due to these attributes, labile organic carbon (LOC) has been identified as an indicator parameter for soil health. Quantifying the turnover rate of LOC also aids in understanding changes in soil nutrient cycling processes. A sequential fumigation incubation method has been developed to estimate soil LOC and potential C turnover rate. The method requires fumigating soil samples and quantifying CO2-C respired during a 10 day incubation period over a series of fumigation-incubation cycles. Labile organic C and potential C turnover rate are then extrapolated from accumulated CO2 with a negative exponential model. Procedures for conducting this method are described.

Labile organic carbon (LOC) and the potential carbon turnover rate are sensitive indicators of changes in soil nutrient cycling processes. Details are provided for a method based on fumigating and incubating soil in a series of cycles and using the CO2 accumulated during the incubation periods to estimate these parameters.

Management practices and environmental changes can alter soil nutrient and carbon cycling. Soil labile organic carbon, a readily decomposable C pool, is ...

Evaluating the Effect of Environmental Chemicals on Honey Bee Development from the Individual to Colony Level

The presence of pesticides in the beekeeping environment is one of the most serious problems that impacts the life of a honey bee. Pesticides can be brought back to the beehive after the bees have foraged on flowers that have been sprayed with pesticides. Pesticide contaminated food can be exchanged between workers which then feed larvae and therefore can potentially affect the development of honey bees. Thus, residual pesticides in the environment can become a chronic damaging factor to honey bee populations and gradually lead to colony collapse. In the presented protocol, honey bee feeding methods are described and applied to either an individual honey bee or to a colony. Here, the insect growth regulator (IGR) pyriproxyfen (PPN), which is widely used to control pest insects and is harmful to the development of honey bee larvae and pupae, is used as the pesticide. The presenting procedure can be applied to other potentially harmful chemicals or honeybee pathogens for further studies.

Herein we present a method to feed pesticide contaminated food to both an individual honey bee and a beehive colony. The procedure evaluates the pesticide effect on individual honey bees by in vivo feeding of basic larval diet and also on the natural condition of beehive colony.

The presence of pesticides in the beekeeping environment is one of the most serious problems that impacts the life of a honey bee. Pesticides can be ...

Self-standing Electrochemical Set-up to Enrich Anode-respiring Bacteria On-site

Anaerobic respiration coupled with electron transport to insoluble minerals (referred to as extracellular electron transport [EET]) is thought to be critical for microbial energy production and persistence in many subsurface environments, especially those lacking soluble terminal electron acceptors. While EET-capable microbes have been successfully isolated from various environments, the diversity of bacteria capable of EET is still poorly understood, especially in difficult-to-sample, low energy or extreme environments, such as many subsurface ecosystems. Here, we describe an on-site electrochemical system to enrich EET-capable bacteria using an anode as a respiratory terminal electron acceptor. This anode is connected&#160;to a cathode capable of&#160;catalyzing abiotic oxygen reduction. Comparing this approach with electrocultivation methods that use a potentiostat for poising the electrode potential, the two-electrode system does not require an external power source. We present an example of our on-site enrichment utilized in an alkaline pond at the Cedars, a terrestrial serpentinization site in Northern California. Prior attempts to cultivate mineral reducing bacteria were unsuccessful, which is likely due to the low-biomass nature of this site and/or the low relative abundance of metal reducing microbes. Prior to implementing our two-electrode enrichment, we measured the vertical profile of dissolved oxygen concentration. This allowed us to place the carbon felt anode and platinum-electroplated carbon felt cathode at depths that would support anaerobic and aerobic processes, respectively. Following on-site incubation, we further enriched the anodic electrode in the laboratory and confirmed&#160;a distinct microbial community compared to the surface-attached or biofilm communities normally observed at the Cedars. This enrichment subsequently led to the isolation of the first electrogenic microbe from the Cedars. This method of on-site microbial enrichment has the potential to greatly enhance the isolation of EET-capable bacteria from low biomass or difficult to sample habitats.

On-site microbial enrichment or in situ cultivation techniques can facilitate the isolation of difficult-to-culture microbial taxa, especially from low-biomass or geochemically extreme environments. Here, we describe an electrochemical set-up without using an external power source to enrich microbial strains that are capable of extracellular electron transport (EET).

Anaerobic respiration coupled with electron transport to insoluble minerals (referred to as extracellular electron transport [EET]) is thought to be ...

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 ...

Composition and Distribution Analysis of Bioaerosols Under Different Environmental Conditions

Variable microorganisms in particulate matter (PM) under different environmental conditions may have significant impacts on human health. In this study, we described a protocol for multiple analyses of the biological compositions in environmental PM. Five experiments are presented: (1) PM number monitoring by using a laser particle counter; (2) PM collection by using a cyclonic aerosol sampler; (3) PM collection by using a high-volume air sampler with filters; (4) culturable microbes collection by the Andersen six-stage sampler; and (5) detection of biological composition of environmental PM by bacterial 16SrDNA and fungal ITS region sequencing. We selected hazy days and a livestock farm as two typical examples of the application in this protocol. In this study, these two sampling methods, cyclonic aerosol sampler and filter sampler, showed different sampling efficiency. The cyclonic aerosol sampler performed much better in terms of collecting bacteria, while these two methods showed the same efficiency in collecting fungi. Filter samplers can work under low temperature conditions while cyclonic aerosol samplers have a sampling limitation for temperature. A solid impacting sampler, such as an Andersen six-stage sampler, can be used to sample bioaerosols directly into the culture medium, which increases the survival rate of culturable microorganisms. However, this method mainly relies on culture while more than 99% of microbes cannot be cultured. DNA extracted from the culturable bacteria collected by the Andersen six-stage sampler and samples collected by cyclonic aerosol sampler and filter sampler were detected by bacterial 16S rDNA and fungal ITS region sequencing.All the methods above may have wide application in many fields of study, such as environmental monitoring and airborne pathogen detection. From these results, we can conclude that these methods can be used under different conditions and may help other researchers further explore the health impacts of environmental bioaerosols.

Understanding the biological composition of environmental particulate matter is important for the study of its significant impacts on human health and disease spread. Here, we used three types of bioaerosol sampling methods and a biological analysis of airborne microbes to better explore airborne microbial communities under different environmental conditions.

Variable microorganisms in particulate matter (PM) under different environmental conditions may have significant impacts on human health. In this study, ...

PARbars: Cheap, Easy to Build Ceptometers for Continuous Measurement of Light Interception in Plant Canopies

Ceptometry is a technique used to measure the transmittance of photosynthetically active radiation through a plant canopy using multiple light sensors connected in parallel on a long bar. Ceptometry is often used to infer properties of canopy structure and light interception, notably leaf area index (LAI) and effective plant area index (PAIeff). Due to the high cost of commercially available ceptometers, the number of measurements that can be taken is often limited in space and time. This limits the usefulness of ceptometry for studying genetic variability in light interception, and precludes thorough analysis of, and correction for, biases that can skew measurements depending on the time of day. We developed continuously logging ceptometers (called PARbars) that can be produced for USD $75 each and yield high quality data comparable to commercially available alternatives. Here we provide detailed instruction on how to build and calibrate PARbars, how to deploy them in the field and how to estimate PAI from collected transmittance data. We provide representative results from wheat canopies and discuss further considerations that should be made when using PARbars.

Here, we present detailed instructions on how to build and calibrate research quality ceptometers (light sensors that integrate light intensity across many sensors arrayed linearly along a horizontal bar).

Ceptometry is a technique used to measure the transmittance of photosynthetically active radiation through a plant canopy using multiple light sensors ...

Quantification of the Potential Impact of Glyphosate-Based Products on Microbiomes

Glyphosate-based products (GBP) are the most common broad-spectrum herbicides worldwide. The target of glyphosate is the enzyme 5-enolpyruvylshikimate-3-phosphate synthase (EPSPS) in the shikimate pathway, which is virtually universal in plants. The inhibition of the enzyme stops the production of three essential amino acids: phenylalanine, tyrosine, and tryptophan. EPSPS is also present in fungi and prokaryotes, such as archaea and bacteria; thus, the use of GBP may have an impact on the microbiome composition of soils, plants, herbivores, and secondary consumers. This article aims to present general guidelines to assess the effect of GBP on microbiomes from field experiments to bioinformatics analyses and provide a few testable hypotheses. Two field experiments are presented to test the GBP on non-target organisms. First, plant-associated microbes from 10 replicated control and GBP treatment plots simulating no-till cropping are sampled and analyzed. In the second experiment, samples from experimental plots fertilized by either poultry manure containing glyphosate residues or non-treated control manure were obtained. Bioinformatics analysis of EPSPS protein sequences is utilized to determine the potential sensitivity of microbes to glyphosate. The first step in estimating the effect of GBP on microbiomes is to determine their potential sensitivity to the target enzyme (EPSPS). Microbial sequences can be obtained either from public repositories or by means of PCR amplification. However, in the majority of field studies, microbiome composition has been determined based on universal DNA markers such as the 16S rRNA and the internal transcribed spacer (ITS). In these cases, sensitivity to glyphosate can only be estimated through a probabilistic analysis of EPSPS sequences using closely related species. The quantification of the potential sensitivity of organisms to glyphosate, based on the EPSPS enzyme, provides a robust approach for further experiments to study target and non-target resistant mechanisms.

Glyphosate-based products (GBP) are the most common broad-spectrum herbicides worldwide. In this article, we introduce general guidelines to quantify the effect of GBP on microbiomes, from field experiments to bioinformatics analyses.