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

Megumi Kuroiwa; Keitaro Fukushima; Kazuma Hashimoto; Yukiko Senga; Tsubasa Sato; Chie Katsuyama; Yuichi Suwa

doi:10.3791/59562

JoVE Journal > Environment

Environment

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

Published: October 07, 2020

doi:

10.3791/59562

Megumi Kuroiwa¹, Keitaro Fukushima^2,3, Kazuma Hashimoto², Yukiko Senga⁴, Tsubasa Sato⁴, Chie Katsuyama⁵, Yuichi Suwa¹

¹Department of Biological Sciences, Faculty of Science and Engineering,Chuo University, ²Faculty & Graduate School of Urban Environmental Sciences,Tokyo Metropolitan University, ³Center for Ecological Research,Kyoto University, ⁴Department of Chemistry, Faculty of Science,Toho University, ⁵Graduate School of Integrated Arts and Sciences,Hiroshima University

Summary

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.

Abstract

The importance of understanding the fate of nitrate (NO₃⁻), 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⁻ 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⁻ 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⁻ 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⁻ via persulfate oxidation. Third, the NO₃⁻ is converted to N₂O via an N₂O reductase deficient denitrifier. Finally, the converted N₂O is analyzed using a previously developed quadrupole gas chromatography–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.

Introduction

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 NO₃⁻². This may cause environmental problems such as drinking water pollution, eutrophication, and the formation of hypoxia. NO₃⁻ 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⁻. Denitrification is the microbial reduction of NO₃⁻ 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⁻ 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⁻ 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⁻ from terrestrial ecosystems, and in previous studies DNRA has been shown to contribute over a very wide range of NO₃⁻ removal (0–99%)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⁻ 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⁻ as a tracer.¹⁵N-labeled NO₃⁻ 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⁻, (2) extraction and recovery of NH₄⁺ using a “diffusion procedure” with modifications, (3) persulfate oxidation of NH₄⁺ in the sample, consisting of indigenous NH₄⁺ and ¹⁵NH₄⁺ derived from ¹⁵NO₃⁻ via DNRA activity, into NO₃⁻ and ¹⁵NO₃⁻, (4) subsequent microbial transformation of NO₃⁻ and ¹⁵NO₃⁻ to N₂O isotopomers via the modified denitrifier method, and (5) quantification of the N₂O isotopomers using gas chromatography–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.

Protocol

1. Preparation of a PTFE envelope for quantitatively capturing gaseous NH₃

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).
Ash a glass fiber filter (10 mm in diameter with a pore size of 2.7 μm) at 450 °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).
Spot 20 µL 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 1.4–1.7, are shown in Figure 1 and should be conducted swiftly.
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.
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.
Fold the open end with the tweezers, and then press the edge with the tweezers (Figure 1d).
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.

2. Preparing the biomass of a nitrous oxide reductase deficient denitrifier, Pseudomonas chlororaphis subsp. aureofaciens ATCC13985, for the denitrifier method

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 °C for 2–3 days.
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 °C in dark until obtaining maximum growth; this will be used as the preculture.
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 °C. After cultivating for 8 h, replace the silicon rubber stopper with a screw cap and close tightly. Continue the cultivation in anoxia overnight.
Centrifuge the culture at 18,800 x g for 15 min at 4 °C to obtain biomass pellets.
Wash the packed biomass three times with 30 mL of Dulbecco’s phosphate-buffered saline (D-PBS(-), pH 7.5), to completely eliminate the NO₃⁻. The conditions of the centrifugation are same as in Step 2.3.
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 −80 °C until use (see section 6).

3. Elimination of oxygen, nitrite, and nitrate from the sample sediment

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).
Seal the vial with a black butyl rubber stopper (washed with ion-exchanged water and sterilized via autoclaving) and an aluminum cap.
Purge the suspension and replace the headspace air with Ar (>99.99%) at 0.6 L/min for 20 min using a manifold.
Replace the Ar headspace gas with ultra-pure (>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.
Incubate the vials at 20 °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.

4. Time course experiment for determining DNRA rate

Replace the headspace gas with fresh ultra-pure He using the same procedure as in step 3.5 but without the four repetitions.
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.
Incubate vials at 20 °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 4.4–4.9.
Remove the aluminum cap and butyl rubber stopper from each vial. Then, add KCl (ashed at 450 °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.
Shake the sediment at 150 rpm for 1 h at 4 °C under dark conditions to extract the NH₄⁺.
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 °C.
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 µm, 25 mm in diameter). Then, rinse the membrane filter with 1 mL of supernatant. Place the rinsed syringe–filter unit onto a 20-mL wide-mouth polypropylene (PP) bottle.
Filter the remaining supernatant through an acetate membrane filter into the 20-mL PP bottle. Store the extracts extracts at −20 °C until further analysis.

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⁻

Prepare standard solutions of ¹⁴NH₄Cl with concentration gradients of 0 μmol/L, 10 μmol/L, 40 μmol/L, 100 μmol/L, 200 μmol/L, 400 μmol/L, and 500 μmol/L. For the ¹⁵N ratio analysis, fix the total concentration of NH₄⁺ to 200 μmol/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.
Transfer 30 mg of MgO (ashed at 450 °C for 4 h) to a 20-mL glass vial, and place the PTFE envelope in the vial.
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 μmol/L, dilute the sample to below 500 μmol/L.
Shake the vials at 150 rpm for 3 h at 4 °C under dark conditions.
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.
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 1.4–1.7.
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.
Repeat steps 5.5–5.7 for the remaining envelopes.
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.
Prepare the persulfate-oxidizing solution (POR) reagent²⁵^,²⁶.
1. Because it cannot be stored, prepare the exact amount of POR needed for treating a single day of samples.
2. 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.
3. 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⁻ because tap water normally contains NO₃⁻.
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.
Stand the test tubes on a rack, wrap them in double-layered aluminum foil, and autoclave them for 1 h at 121 °C. Keep the tubes in an upright position during this step and avoid rapid changes in temperature after finishing the autoclaving.

6. Determining the NO₃⁻ converted from NH₄⁺ by the denitrifier method using quadrupole GC/MS

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).
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 (>99.995%) stream for 1 h.
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.
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.
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.
Incubate the vials overnight at 25 °C under dark conditions.
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.
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.

7. Data analysis

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⁺.
Calculate the potential DNRA rate using equations provided elsewhere²⁸^,²⁹^,³⁰.

Representative Results

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 (0–3 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 °C until the DNRA analysis was conducted.

The incubation procedures for the ¹⁵NO₃⁻ 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⁻ pool²⁹. The calculated rates were within the range of 24.8–177 nmol-N g⁻¹ dry soil h⁻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).

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)–(E). The acidified filter inside the envelope captures the gaseous NH₃. These steps should be conducted quickly. Detailed information is given in steps 1.2–1.7 in the protocol section. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

	¹⁵N-labeled and non-labeled substrates added to each vial
	100mM	100mM
	NH₄Cl	K¹⁵NO₃
Volume (µL) of 100 mM stock solution added to each vial	24	60
Final concentration (µmol/L)*	>230^§	570
*shown values are calculated by assuming that water content of the sediment is 50%
§depending on the background ammonium concentration

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.

Equipment
quadrupole GCMS	Shimadzu GCMS-QP2010 Ultra
column	CP-PoraBONDQ 25m; φ 0.32mm; film thickness, 5µm
Analytical conditions
column temp	40 °C
injection port temp	100 °C
carrier gas stream	Total flow rate, 47.1 mL•min^-1 flow rate in column, 2.10 mL•min^-1
sprit ratio	20
detection voltage	1.5 kv
Sensitivity of N₂O
lower limit of detection (LOD)*	1.42 pmol
lower limit of quantification (LOQ)*	4.58 pmol
*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.

Table 2: Conditions for the GC/MS analysis.

sediment	DNRA rate	enrichment of NO₃^–*
	nmol-N g^-1 hr^-1	atom%
Intertidal 1	177	99.9
Intertidal 2	129	99.0
Subtidal 1	39.3	99.9
Subtidal 2	24.8	99.0
*same as atom% of added KNO₃; complete elimination and no nitrification under used incubation conditions was tested previously.

Table 3: Potential DNRA rates of the tested intertidal and subtidal sediments.

Discussion

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 N₂via 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 pK_a, 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 μmol and 0.09 μmol, respectively, and these values are equivalent to 6 μmol/L and 18 μmol/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.

Disclosures

The authors have nothing to disclose.

Acknowledgements

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

Materials

¹⁵N-KNO₃	SHOKO SCIENCE	N15-0197
¹⁵N-NH₄Cl	SHOKO SCIENCE	N15-0034
20 mL PP bottle	SANPLATEC	61-3210-18	Wide-mouth
Aluminum cap	Maruemu	1307-13	No. 20, with hole
Boric acid	Wako	021-02195
Centrifuge	HITACHI	Himac CR21G II
Deoxygenized Gas Pressure & Replace Injector	SANSIN INDUSTRIAL	IP-12
Disposable cellulose acetate membrane filter	ADVANTEC	25CS020AS	Pore size 0.22 µm, 25 mm in diameter
Disposable syringe	Termo	SS-10SZ	10 mL
Disposable syringe	Termo	SS-01T	1 mL
Dulbecco’s Phosphate Buffered Saline (-)	NISSUI PHARMACEUTICAL	5913
Gastight syringe	VICI Valco Instruments	4075-15010	Series A-2, 100 µL
GC/MS	shimadzu	GCMS-QP2010ultra
GF/D	Whatman	1823-010	10 mm in diameter
Glass vial	Maruemu	0501-06	20 mL
Gray butyl rubber stopper	Maruemu	1306-03	No.20-S
H₂SO₄	Wako	192-04696	Guaranteed Reagent
K₂S₂O₈	Wako	169-11891	Nitrogen and Phosphorus analysis grade
KCl	Wako	163-03545	Guaranteed Reagent
KNO₃	Wako	160-04035	Guaranteed Reagent
NaOH	Wako	191-08625	Nitrogen compounds analysis grade
NH₄Cl	Wako	017-02995	Guaranteed Reagent
Plastic centrifuge tube	ASONE	1-3500-22	50 mL, VIO-50BN
Pseudomonas chlororaphis subsp. aureofaciens	American Type Culture Collection (ATCC)	ATCC 13985	Freeze-dried, the type strain of Pseudomonas aureofaciens
PTFE sealing tape	Sigma-Aldrich	Z221880	25 mm in width
Reciprocating shaker	TAITEC	0000207-000	NR-10
Screw-cap test tube	IWAKI	84-0252	11 mL
PTFE-lined cap for test tube	IWAKI	84-0262
Tryptic Soy Broth	Difco Laboratories	211825

References

Gruber, N., Galloway, J. N. An Earth-system perspective of the global nitrogen cycle. Nature. 451 (7176), 293-296 (2008).
Galloway, J. N., et al. The Nitrogen Cascade. Bioscience. 53 (4), 341-356 (2003).
Rysgaard, S., Risgaard-Petersen, N., Sloth, N. P., Caumette, P., Castel, J., Herbert, R. Nitrification, denitrification, and nitrate ammonification in sediments of two coastal lagoons in Southern France. Coastal Lagoon Eutrophication and Anaerobic Processes (C.L.E.AN.). Developments in Hydrobiology. 117, 133-141 (1996).
Christensen, P. B., Rysgaard, S., Sloth, N. P., Dalsgaard, T., Schwærter, S. Sediment mineralization, nutrient fluxes, denitrification and dissimilatory nitrate reduction to ammonium in an estuarine fjord with sea cage trout farms. Aquatic Microbial Ecology. 21, 73-84 (2000).
Tobias, C. R., Anderson, I. C., Canuel, E. A., Macko, S. A. Nitrogen cycling through a fringing marsh-aquifer ecotone. Marine Ecology Progress Series. 210, 25-39 (2001).
An, S. M., Gardner, W. S. Dissimilatory nitrate reduction to ammonium (DNRA) as a nitrogen link, versus denitrification as a sink in a shallow estuary (Laguna Madre/Baffin Bay, Texas). Marine Ecology Progress Series. 237, 41-50 (2002).
Gardner, W. S., et al. Nitrogen fixation and dissimilatory nitrate reduction to ammonium (DNRA) support nitrogen dynamics in Texas estuaries. Limnology & Oceanography. 51 (1), 558-568 (2006).
Preisler, A., et al. Biological and chemical sulfide oxidation in a Beggiatoa inhabited marine sediment. The ISME Journal. 1 (4), 341-353 (2007).
Gardner, W. S., McCarthy, M. J. Nitrogen dynamics at the sediment-water interface in shallow, sub-tropical Florida Bay: why denitrification efficiency may decrease with increased eutrophication. Biogeochemistry. 95 (2-3), 185-198 (2009).
Dong, L. F., et al. Changes in benthic denitrification, nitrate ammonification, and anammox process rates and nitrate and nitrite reductase gene abundances along an estuarine nutrient gradient (the Colne estuary, United Kingdom). Applied and Environmental Microbiology. 75 (10), 3171-3179 (2009).
Koop-Jakobsen, K., Giblin, A. E. The effect of increased nitrate loading on nitrate reduction via denitrification and DNRA in salt marsh sediments. Limnology & Oceanography. 55 (2), 789-802 (2010).
Dong, L. F., et al. Dissimilatory reduction of nitrate to ammonium, not denitrification or anammox, dominates benthic nitrate reduction in tropical estuaries. Limnology & Oceanography. 56 (1), 279-291 (2011).
Fernandes, S. O., Bonin, P. C., Michotey, V. D., Garcia, N., LokaBharathi, P. A. Nitrogen-limited mangrove ecosystems conserve N through dissimilatory nitrate reduction to ammonium. Scientific Reports. 2, 419 (2012).
Behrendt, A., de Beer, D., Stief, P. Vertical activity distribution of dissimilatory nitrate reduction in coastal marine sediments. Biogeosciences. 10 (11), 7509-7523 (2013).
Song, G. D., Liu, S. M., Marchant, H., Kuypers, M. M. M., Lavik, G. Anammox denitrification and dissimilatory nitrate reduction to ammonium in the East China Sea sediment. Biogeosciences. 10 (11), 6851-6864 (2013).
Yin, G., Hou, L., Liu, M., Liu, Z., Gardner, W. S. A novel membrane inlet mass spectrometer method to measure 15NH4+15+ for isotope-enrichment experiments in aquatic ecosystems. Environmental Science & Technology. 48 (16), 9555-9562 (2014).
Zheng, Y., et al. Tidal pumping facilitates dissimilatory nitrate reduction in intertidal marshes. Scientific Reports. 6, 21338 (2016).
Bu, C., et al. Dissimilatory Nitrate Reduction to Ammonium in the Yellow River Estuary: Rates, Abundance, and Community Diversity. Scientific Reports. 7, 6830 (2017).
Scott, J. T., McCarthy, M. J., Gardner, W. S., Doyle, R. D. Denitrification, dissimilatory nitrate reduction to ammonium, and nitrogen fixation along a nitrate concentration gradient in a created freshwater wetland. Biogeochemistry. 87 (1), 99-111 (2008).
Shan, J., et al. Dissimilatory Nitrate Reduction Processes in Typical Chinese Paddy Soils: Rates, Relative Contributions, and Influencing Factors. Environmental Science & Technology. 50 (18), 9972-9980 (2016).
Silver, W. L., Herman, D. J., Firestone, M. K. Dissimilatory nitrate reduction to ammonium in upland tropical forest soils. Ecology. 82 (9), 2410-2416 (2001).
Holmes, R. M., McClelland, J. W., Sigman, D. M., Fry, B., Peterson, B. J. Measuring 15N–NH4+ in marine, estuarine and fresh waters: An adaption of the ammonia diffusion method for samples with low ammonium concentrations. Marine Chemistry. 60 (3-4), 235-243 (1998).
Sigman, D. M., et al. A bacterial method for the nitrogen isotopic analysis of nitrate in seawater and freshwater. Analytical Chemistry. 73 (17), 4145-4153 (2001).
Weigand, M. A., Foriel, J., Barnett, B., Oleynik, S., Sigman, D. M. Updates to instrumentation and protocols for isotopic analysis of nitrate by the denitrifier method. Rapid Communications in Mass Spectrometry. 30 (12), 1365-1383 (2016).
Isobe, K., et al. Analytical techniques for quantifying 15N/14N of nitrate, nitrite, total dissolved nitrogen and ammonium in environmental samples using a gas chromatograph equipped with a quadrupole mass spectrometer. Microbes and Environments. 26 (1), 46-53 (2011).
Miyajima, T., Tanaka, Y., Koile, Y. Determining 15N enrichment of dissolved organic nitrogen in environmental waters by gas chromatography/negative-ion chemical ionization mass spectrometry. Limnology and Oceanography. 3 (3), 164-173 (2005).
Stevens, R. J., Laughlin, R. J., Burns, L. C., Arah, J. R. M., Hood, R. C. Measuring the contributions of nitrification and denitrification to the flux of nitrous oxide from soil. Soil Biology and Biochemistry. 29 (2), 139-151 (1997).
Porubsky, W. P., Velasquez, L. E., Joye, S. B. Nutrient-replete benthic microalgae as a source of dissolved organic carbon to coastal waters. Estuaries and Coasts. 31 (5), 860-876 (2008).
Huygens, D., et al. Mechanisms for retention of bioavailable nitrogen in volcanic rainforest soils. Nature Geoscience. 1 (8), 543-548 (2008).
Rutting, T., Boeckx, P., Muller, C., Klemedtsson, L. Assessment of the importance of dissimilatory nitrate reduction to ammonium for the terrestrial nitrogen cycle. Biogeosciences. 8 (7), 1779-1791 (2011).
Song, B., Lisa, J. A., Tobias, C. R. Linking DNRA community structure and activity in a shallow lagoonal estuarine system. Frontiers in Microbiology. 5, 460 (2014).
Cheng, L., et al. Dissimilatory nitrate reduction processes in sediments of urban river networks: Spatiotemporal variations and environmental implications. Environmental Pollution. 219, 545-554 (2016).
Lisa, J. A., Song, B., Tobias, C. R., Hines, D. E. Genetic and biogeochemical investigation of sedimentary nitrogen cycling communities responding to tidal and seasonal dynamics in Cape Fear River Estuary. Estuarine, Coastal and Shelf Science. 167, A313-A323 (2015).
Deng, F. Y., et al. Dissimilatory nitrate reduction processes and associated contribution to nitrogen removal in sediments of the Yangtze Estuary. Journal of Geophysical Research: Biogeosciences. 120 (8), 1521-1531 (2015).
Tiedje, J. M., Zehnder, A. J. B. . Biology of Anaerobic Microorganisms. , 179-244 (1988).
Tiedje, J. M., Sexstone, A. J., Myrold, D. D., Robinson, J. A. Denitrification: ecological niches, competition and survival. Antonie van Leeuwenhoek. 48, 569-583 (1982).
Hardison, A. K., Algar, C. K., Giblin, A. E., Rich, J. J. Influence of organic carbon and nitrate loading on partitioning between dissimilatory nitrate reduction to ammonium (DNRA) and N2 production. Geochimica et Cosmochimica Acta. , 164 (2015).
Sigman, D. M., et al. Natural abundance-level measurement of the nitrogen isotopic composition of oceanic nitrate: an adaptation of the ammonia diffusion method. Marine Chemistry. 57 (3-4), 227-242 (1997).
Risgaard-Petersen, N., Rysgaard, S., Revsbech, N. P. Combined microdiffusion-hypobromite oxidation method for determining nitrogen-15 isotope in ammonium. Soil Science Society of America Journal. 59 (4), (1995).
Gardner, W. S., Bootsma, H. A., Evans, C., John, P. A. S. Improved chromatographic analysis of 15N:14N ratios in ammonium or nitrate for isotope addition experiments. Marine Chemistry. 48 (3-4), 271-282 (1995).

Play Video

PDF

DOI

DOWNLOAD MATERIALS LIST

Cite This Article

Kuroiwa, M., Fukushima, K., Hashimoto, K., Senga, Y., Sato, T., Katsuyama, C., Suwa, Y. Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on ¹⁴NH₄⁺/¹⁵NH₄⁺ Analyses via Sequential Conversion to N₂O. J. Vis. Exp. (164), e59562, doi:10.3791/59562 (2020).

Automatically Generated