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.
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.
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.
1. Preparation of a PTFE envelope for quantitatively capturing gaseous NH3
2. Preparing the biomass of a nitrous oxide reductase deficient denitrifier, Pseudomonas chlororaphis subsp. aureofaciens ATCC13985, for the denitrifier method
3. Elimination of oxygen, nitrite, and nitrate from the sample sediment
4. Time course experiment for determining DNRA rate
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−
6. Determining the NO3− converted from NH4+ by the denitrifier method using quadrupole GC/MS
7. Data analysis
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.
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 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.
The authors have nothing to disclose.
We thank Naoto Tanaka for helping data collection and developing the protocol. The collection of samples was supported by JSPS KAKENHI Grant Number 17K15286.
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 |