The Potential Inhibitory Effect of Copper and Nickel on Gaseous Soil N Emissions

Abstract

Nickel (Ni) and copper (Cu) are essential soil micronutrients that are beneficial for plant growth and development when applied at the right levels to deficient soils. However, they can also be of concern if excessive application leads to accumulation in soil, causing soil fertility problems that can compromise food production as well as result in human exposure to these metals if they enter the food chain. The application of Ni and Cu salts can strongly influence the nitrogen (N) cycle as they can stimulate soil microbiological activity that leads to N losses to the atmosphere. A laboratory incubation was conducted to study the effect of these metal salts on gaseous N emissions. Copper sulphate (CuSO₄) and nickel sulphate (NiSO₄) were applied at rates below soil toxicity (3.0 kg Ni/ha and 7.5 kg Cu/ha), together with ¹⁵N-labelled-urea (rate 100 kg N/ha) and under two soil water-filled pore space (WFPS) conditions: continuous anaerobic (75% WFPS) versus aerobic (50% WFPS) followed by an anaerobic (75% WFPS) cycle. Results showed no significant effect of the Ni application on nitric oxide (NO) emissions at the low rates applied. However, there was an indication of a meaningful (p = 0.051) production of this gas under aerobic soil conditions. No N₂ emissions were observed from any treatment (only expected to occur under anaerobic conditions) during the 43 days of the incubation, which could suggest that, as for Ni, the selected Cu application rate may not clearly stimulate the complete denitrification process to produce N₂. It is possible that the expected increase in N₂ emissions or larger NO emissions were counteracted by the low amount of soil-extractable Cu and Ni available after application because of binding processes with the clay fraction (48% clay) and the acid nature of the Cambisol used in the incubation. Further research into the effect of metal applications to contrasting soil types, with varied application rates and repeated application events, and under differing soil moisture conditions on N emissions is required. Furthermore, experiments should also be conducted at finer scales to elucidate the specific effect of these metal application rates on the biochemical processes responsible for the N gases produced.

1. Introduction

Nitrogen (N) losses from agricultural soils in the form of leaching and in the form of gaseous emissions are an environmental concern. Losses to the atmosphere mainly occur via gaseous emissions of ammonia (NH₃), nitric oxide (NO), nitrous oxide (N₂O) and nitrogen (N₂). Plants need N in its inorganic form for uptake as do the microorganisms in the soil involved in the processes that release these N gases. One important process in the N cycle that contributes to the pool of inorganic N is mineralisation, where organic biomass is transformed into soluble forms (inorganic) by microbes and soil organisms as their source of carbon (C) and N. Mineral forms of N are released into the soil once these organisms meet their needs, and these excess inorganic forms are available for further use by plant or can be lost to the environment. However, if microbial demand for N is high and/or N sources are not enough, microbes retain or immobilise N in their biomass [1]. Applications of organic and mineral fertilisers to agricultural soils are often conducted by farmers to provide plants with the required amount of N for development and growth. However, these activities can also lead to great losses of N to the environment, causing not just serious environmental problems but also an economic loss for farmers. Losses via NH₃ mainly occur through volatilisation of urea contained in animal faeces and fertilisers like straight urea, while NO, as well as N₂O, are mainly produced during nitrification and denitrification processes of the soil N cycle [2]. When using chemical fertilisers containing urea CO(NH₂)₂ salts, the added urea is hydrolysed into ammonia (NH₃) with the help of urease. In an acidic environment, NH₃ is dissolved and converted into NH₄⁺, which is available to plants and subjected to further nitrification in aerobic conditions by soil nitrifiers, including nitrifying bacteria, fungi and archaea [1]. NH₃ is oxidised into hydroxylamine (NH₂OH), and this further oxidised to nitrite (NO₂⁻), forming NO as an intermediary compound. This gas can also be produced (under low O₂ levels), together with N₂O, during the dissimilatory reduction of NO₂⁻, nitrate (NO₃⁻) or both [3]. This gas has a negative impact on the environment, as it is a highly reactive gas reacting with volatile organic compounds and sunlight, increasing ground level ozone, while acting as a catalyst in the stratosphere and breaking down the ozone layer [4]. It also contributes to acid rain when it oxidises in the air to form NO₂, which then reacts with water vapour to form nitric acid [5]. It has been estimated that soils contribute annually 3.3 to 21 Tg N ha⁻¹ to NO emissions [6,7], which highlights the importance of mitigating these emissions to achieve better soil N management and reduce environmental emissions.

Nitrous oxide is one of the most important greenhouse gases, with a global warming potential 265 times that of carbon dioxide (CO₂) [8] and is known to be responsible for approximately 60% of the total global anthropogenic N (~6 Tg N₂O-N) [9]. With the increasing use of N fertilisers to support increases in food production [10], it is estimated that the emissions of N₂O from agricultural lands will also increase [11,12]. Nitrous oxide is a subproduct of nitrification but it is also produced by denitrification when NO₃⁻ is used by denitrifiers as a terminal electron acceptor instead of O₂ for respiration when O₂ is unavailable [1]. This process produces N₂, as well as N₂O and NO, which is the ultimate end of the denitrification process and, despite not being of environmental concern, it represents an economic loss of the N applied to increase yields.

When looking at the biogeochemical processes that cause NO, N₂O and N₂ emissions, there are many factors that influence and lead to these atmospheric losses of N and soil water content is one of them [13]. The amount of soil water determines the degree of aerobicity and anaerobicity of the soil environment by changing oxygen (O₂) availability for plant roots and microorganisms. These changes have a direct impact on soil microbiology, making soil N biochemical processes highly dynamic and changeable within the soil environment [14,15]. When soil moisture conditions are high, the proportion of soil pores filled with air decrease as they get replaced with water, reducing O₂ diffusivity within the soil [16]. It can also make C and N more available after cycles of soil rewetting after periods of drought. By filling the soil pores and preventing O₂ supply, high moisture content provides suitable conditions for denitrification. Many studies have shown a significant relationship between the water-filled pore space (WFPS) and microbial pathways that produce N gases. It is generally considered [17,18,19] that nitrification is more important than denitrification when WFPS < 60% (more NO than N₂O is produced), denitrification dominates when WFPS > 60% (more N₂O production over NO) and N₂ is mainly produced under total anaerobic soil conditions [18,19]. However, these ranges may vary depending on the type of soil.

Many microbial enzymes involved in the reactions that produce these gases are also dependent on metal cofactors. For example, urease uses a redox-active nickel (Ni) cofactor for the hydrolysis of urea into NH₃ [20,21,22] or nitrous oxide reductase uses copper (Cu) as a redox-active cofactor for the catalysis of the final step of bacterial denitrification to convert N₂O into N₂ and water [3,23]. Thus, the abundance and availability of these metals in the soil is also a key factor as they can influence the soil N cycle and exert control on the resulting N emissions [24]. Metal applications in excess have been reported to have a positive influence on microbial populations and therefore affect processes in the N cycle; for example, they can affect the population of denitrifiers, leading to decreases in N₂O emissions [25]. This could be seen as a positive result, but the application of these metals to soils does not only play an essential part in the microbial N cycle. It is also essential to plants, and if metal applications exceed plant requirements, they can accumulate in the soil or even cause toxicity and the inhibition of ecosystem processes. Approximately 20% of Cu in soil is in a soluble form and, therefore, available for plant use, which can cause Cu deficiency in many agricultural soils worldwide [26]. Animal manures, sewage sludge and biosolids usually contain high concentrations of Cu, which can supply crop Cu requirements (2–20 ppm) for cellular metabolic processes. However, at elevated levels, Cu becomes toxic to plants (>20 ppm, leaf analysis), which is of concern when applying those wastes to soil, with excess Cu inhibiting plant photosynthesis, reducing plant productivity and potentially posing a risk to humans due to accumulation in the food chain [26]. Copper sulphate (CuSO₄) is the most common inorganic source of fertiliser, with Cu rates of 1–22 kg/ha needed to correct a soil deficiency [27]. In contrast to Cu, a deficit of Ni in soils is uncommon as plant requirements of this metal are very low (0.1–1.0 ppm), but soil toxicity is common when biosolids, usually with very high concentrations of Ni, are applied as an amendment.

The UK Government’s recommended rates are intended to reduce heavy metal inputs to minimise their accumulation in agricultural soils and reduce the risk of these metals entering the food chain and affecting human health. However, the specific effect of those metal rates on the N cycle, particularly on N gaseous emissions, has not yet been assessed by the UK Government. In this study, we investigated the impact on N emissions of those two heavy metals (Ni and Cu), when applied to soil cores together with urea under two soil water conditions (continuous anaerobic versus aerobic–anaerobic cycle). Application rates were kept below soil and plant toxicity levels, following the UK Government’s permissible application rates for these metals [28]. Using isotope tracing, this study also analyses the transformation of the applied urea-N into soil ¹⁵NH₄ and soil ¹⁵NO₃ and evaluates the ratio of N₂O derived from ¹⁵N-urea. Anaerobic conditions were maintained by keeping the soil moisture content at 75% WFPS, which was the highest moisture that the soil could hold without collapsing and leaching and at which N₂O reduction to N₂ is mainly produced from denitrification and heterotrophic nitrification [29]. Aerobic conditions were achieved by maintaining soil moisture at 50% WFPS.

2. Materials and Methods

2.1. Experimental Design

The current study was carried out at Rothamsted Research (North Wyke, UK). The experiment included four treatments and three replicates for each treatment. The four treatments are shown in Table 1. In the treatment of T1 and T3, Ni was added as nickel sulphate (NiSO₄) at a rate of 3.0 kg Ni ha⁻¹. In the T2 treatment, Cu was applied as copper sulphate (CuSO₄) at a rate of 7.5 kg Cu ha⁻¹. The quantity of the amendment was based on the highest annual permissible application rates permitted by the UK government [28,30]. Urea was added to all the treatments with ¹⁵N-labelled at an enrichment level of 28.706 atom% and an application rate of 100 kg N ha⁻¹.

Table 1. Soil moisture conditions and amendments used for the experimental treatments.

Treatment	Moisture Conditions (% WFPS)			Soil Amendments
	Day 0 to 14	Day 14 to 42	Treatment ID
CK	50	75	50% WFPS–75% WFPS	15Urea-N
T1	50	75	50% WFPS–75% WFPS	Ni + 15Urea-N
T2	50	75	50% WFPS–75% WFPS	Cu + 15Urea-N
T3	75	75	Continuous 75% WFPS	Ni + 15Urea-N

The experimental duration was 43 days. During this time, two separate incubations were run simultaneously with the following aims:

(i)

One incubation was setup in the automated Denitrification Incubation System (DENIS) [31], which has 12 soil incubation chambers able to contain soil cores of 14.2 cm diameter. These were filled with 3 replicates of each of the 4 treatments. The DENIS is an incubation system where a gas mixture continuously flows in and out of the chambers. Each chamber is selected sequentially every 8 min and the gas stream coming from each chamber is diverted to two gas analysers: a gas chromatograph with a helium ionisation detector (HID) for N₂ determination and a chemiluminescence analyser for nitric oxide (NO). The detection limit for N₂ was 0.5 ppm and the linearity of the detector was observed between 1 and 1000 ppm N₂. The NO analyser shows a linear response between 0.5 ppb and 500 ppm [32]. Gas measurements were conducted every 96 min from each chamber throughout the experimental period. Between samplings, the gas from each chamber flows through an extra sample vial before being vented to the atmosphere. The extra sample vials were exchanged twice a day for later ¹⁵N₂O analyses. The incubation was carried out inside a temperature-controlled cabinet containing the chambers at 20 °C.

(ii)

The second incubation consisted of a parallel set of cores (small cores), where 60 soil cores (diameter = 4.5 cm) were prepared and received the same amendment under the same soil moisture conditions for the 4 treatments. In total, there were five replicate small core sets for destructive sampling to trace the soil N transformation via soil N extractions. All the parallel small soil cores were maintained in a light-shielding controlled-environment cabinet (Versatile Environmental Test Chamber MLR-532H, Panasonic©), where the temperature was maintained at 20 °C and the air moisture at 90%.

2.2. Critical Timings During the Experiment

Prior to the start of the experiment soil cores were placed in the system (labelled as Day −2) and flushed through with a gas He:O₂ mixture (80:20) at a flow rate of 30 mL min⁻¹ per vessel to remove N₂ from the soil cores. The gas mixture aims to replace the N₂ in natural air with He to background N₂ levels and allow measurements of N₂, an indicator of denitrification. On Day 0, the flow rate was changed to 12 mL min⁻¹ per vessel and re-directed to flow over the surface of the soil core to start measuring baseline emissions. Afterwards, the treatment solutions (urea with or without Cu or Ni) were added to the soil cores. The total volumes of the treatment solutions were 50 mL for the large and 5 mL for the small cores, allowing the soil moisture content to reach 50% WFPS after the addition. To raise the soil moisture content from 50% to 75% WFPS, additions of 260 mL and of 26 mL deionised water (DIW) for the large and small cores, respectively, were carried out on Day 14. The experiment ended on Day 43.

2.3. Soil Preparation, Packing and Soil Moisture Maintenance

The soil used in this study was obtained from a permanent pasture site at Rothamsted Research, North Wyke (50:46:10 N, 3:54:05 W). The soil order in the World Reference Base system was Cambisol. The soil was taken from the top 20 cm from the surface. After sampling, roots and plant residues were removed by sieving the soil through stainless-steel meshes (mesh pore sizes = 8–10 mm). The sieved soil was left at room temperature for 1–2 days and then passed through another stainless-steel mesh (pore size = 2 mm). The soil was packed into cores straight after the final sieving. The basic properties of the soil are shown in Table 2.

Table 2. Initial soil properties of the soil used for the experiment.

Soil Properties	Results	Analytical Method
Soil texture	Silt Clay	Laser diffraction
Sand %	6%
Silt %	46%
Clay %	48%
Soil organic matter	5.9%	Loss on ignition
Soil pH	5.1	Soil: water suspension (ratio = 1:2.5)
Total Ni	13.2 mg/kg	Acid digestion & ICP-OES analysis
Total Cu	27.8 mg/kg

The sieved soil was packed to a bulk density of 0.88 g cm⁻³, which equates to the bulk density measured in the field’s surface soil. About 140 g and 1393 g (dry weight basis) of the sieved soil were packed to 10 cm height in each of the parallel small cores and DENIS cores, respectively. The soil moisture content was 27.88% (dry basis) at packing. More water was added to the cores to raise the soil moisture contents (in WFPS%) to those stated in Table 1. The pore space (porosity, Vp) of the current soil was calculated following Equation (1).

$${\mathbf{V}}_{\mathbf{p}}={\mathbf{V}}_{\mathbf{c}}\times \left[\mathbf{1}-{\displaystyle \frac{{\mathbf{D}}_{\mathbf{B}}}{{\mathbf{D}}_{\mathbf{p}}}}\right]$$

(1)

where V_c is the total volume of each core (cm³) and the D_B and D_p the bulk density (g cm⁻³) and particle density (g cm⁻³) of the soil, respectively. The particle density of the soil was 2.545 g cm⁻³. The amount of deionised water to add (mL or g) was calculated following Equation (2).

$$\mathrm{amount}\mathrm{of}\mathrm{water}\mathrm{to}\mathrm{add}=(\mathbf{X}\mathbf{\%}\times {\mathbf{V}}_{\mathbf{p}})-{\mathbf{V}}_{\mathbf{s}\mathbf{o}\mathbf{i}\mathbf{l}}\times \mathbf{1}$$

(2)

where the X% stands for the targeted ultimate WFPS%, V_p is the pore space (mL) and the V_soil is the volume of the moisture that was initially in the soil (mL). The total weights of the parallel cores were checked every three to four days. Additional deionised water was added to the small cores to maintain the targeted soil moisture contents if needed.

2.4. Soil Analyses

Soil samples were taken from the parallel cores for the analysis of extractable N (NH₄⁺ and NO₃⁻), the ¹⁵N enrichment in soil extract and soil pH on Days −2, 7, 14, 29 and 43. Mineral N was determined by extracting 100 g of fresh soil with 200 mL of 2M KCl solution. Soil NH₄⁺ and NO₃⁻ contents were determined in soil KCl extracts (1:2 soil:2M KCl ratio). Soil moisture was gravimetrically determined after oven-drying subsamples at 105 °C for 24 h. The soil pH measurement followed the method of 1:1 extraction with deionised water. Soil ¹⁵NH₄⁺ and ¹⁵NO₃⁻ were determined using the reaction of alkaline sodium hypobromite (NaOBr) with NH₄⁻-N (for ¹⁵NH₄⁺ quantification) and the reaction of NO₂⁻ and hydroxylamine (NH₂OH) (for ¹⁵NO₃⁻ quantification), in both cases to form N₂O [33]. The N₂O produced was transferred to Exetainers (Labco Ltd., Lampeter, Wales) and the enriched N₂O (15N₂O) was determined with Isotope Ratio Mass Spectrometry (IRMS).

Initial soil total Cu and Ni contents were determined by acid digestion in aqua regia, giving the total amounts of Cu and Ni present in the soil. On experimental Days 7 and 14, the immediately bioavailable and mobile fractions of Cu and Ni were determined by extraction with 1M ammonium nitrate (NH₄NO₃). Total elements and exchangeable cations were analysed using an Agilent 5900 SVDV Inductively Coupled Plasma—Optical Emission Spectrometer, ICP-OES (Agilent Technologies LDA, Didcot, UK).

2.5. Data Analysis and Statistics

The percentage of the N₂O derived from the ¹⁵N-labelled urea was calculated based on Equation (3).

$$\mathrm{Urea}\text{-}\mathrm{derived}{\mathrm{N}}_2\mathrm{O}(\%)=({}^{\mathbf{15}}\mathbf{N}_{\mathbf{2}}\mathrm{}{\mathbf{O}}_{\mathbf{S}}-{}^{\mathbf{15}}\mathbf{N}_{\mathbf{N}}/{}^{\mathbf{15}}\mathbf{N}_{\mathbf{EN}})\mathbf{\ast }\mathbf{100}$$

(3)

where ¹⁵N₂O_S is the atomic ratio of ¹⁵N in the N₂O gas of the sample (atom%); ¹⁵N_N: the natural abundance atomic ratio of ¹⁵N (=0.3663%); and ¹⁵N_En is the enrichment level of ¹⁵N in the amended urea (=28.7064%).

The contribution of the ¹⁵N-Urea application to the total soil mineral N pool was calculated as follows:

$$\mathrm{N}{\mathrm{H}}_4\mathrm{f}\mathrm{r}\mathrm{o}\mathrm{m}\mathrm{a}\mathrm{m}\mathrm{e}\mathrm{n}\mathrm{d}\mathrm{m}\mathrm{e}\mathrm{n}\mathrm{t}\left(\mathrm{\%}\right)=(\mathrm{s}\mathrm{o}\mathrm{i}\mathrm{l}{}^{15}\mathrm{N}\text{-}\mathrm{N}{\mathrm{H}}_4/\mathrm{t}\mathrm{o}\mathrm{t}\mathrm{a}\mathrm{l}\mathrm{N}\text{-}\mathrm{N}{\mathrm{H}}_4)\mathbf{\ast }100$$

(4)

$$\mathrm{T}\mathrm{O}\mathrm{N}\mathrm{f}\mathrm{r}\mathrm{o}\mathrm{m}\mathrm{a}\mathrm{m}\mathrm{e}\mathrm{n}\mathrm{d}\mathrm{m}\mathrm{e}\mathrm{n}\mathrm{t}\left(\mathrm{\%}\right)=(\mathrm{s}\mathrm{o}\mathrm{i}\mathrm{l}{}^{15}\mathrm{N}\text{-}\mathrm{T}\mathrm{O}\mathrm{N}/\mathrm{t}\mathrm{o}\mathrm{t}\mathrm{a}\mathrm{l}\mathrm{N}\text{-}\mathrm{T}\mathrm{O}\mathrm{N})\mathbf{\ast }100$$

(5)

where soil ¹⁵N-NH₄ and soil ¹⁵N-TON are the amounts of soil ¹⁵NH₄ and ¹⁵TON, respectively, both expressed as mg N/kg soil, and total N-NH₄ and total N-TON are the concentrations of NH₄ and TON in the KCl extracts expressed as mg N/kg soil.

The amount of ¹⁵N-NH₄ and soil ¹⁵N-TON in the soil KCl extracts were calculated as follows:

$${}^{15}\mathrm{N}\text{-}\mathrm{N}{\mathrm{H}}_4(\mathrm{m}\mathrm{g}/\mathrm{k}\mathrm{g})=\left[\right({}^{15}\mathrm{N}\text{-}\mathrm{N}{\mathrm{H}}_4\text{-}{}^{15}\mathrm{N}_{\mathrm{N}})/{}^{15}\mathrm{N}_{\mathrm{EN}}]\mathbf{\ast }\mathrm{t}\mathrm{o}\mathrm{t}\mathrm{a}\mathrm{l}\mathrm{N}\text{-}\mathrm{N}{\mathrm{H}}_4$$

(6)

$${}^{15}\mathrm{N}\text{-}\mathrm{T}\mathrm{O}\mathrm{N}(\mathrm{m}\mathrm{g}/\mathrm{k}\mathrm{g})=\left[\right({}^{15}\mathrm{N}\text{-}\mathrm{T}\mathrm{O}\mathrm{N}\text{-}{}^{15}\mathrm{N}_{\mathrm{N}})/{}^{15}\mathrm{N}_{\mathrm{EN}}]\mathbf{\ast }\mathrm{t}\mathrm{o}\mathrm{t}\mathrm{a}\mathrm{l}\mathrm{N}\text{-}\mathrm{T}\mathrm{O}\mathrm{N}$$

(7)

where ¹⁵N-NH₄ and ¹⁵N-TON are the ¹⁵N abundances of ¹⁵NH₄ and ¹⁵TON in the KCl extracts used for the transformation to N₂O (atom%); ¹⁵N_N: the natural abundance atomic ratio of ¹⁵N of a KCl extract with no soil (=0.3652%); ¹⁵N_En: the enrichment level of ¹⁵N-urea amendment applied (=28.7064%) and total N-NH₄ and total N-TON are the concentrations of NH₄ and TON in the KCl extract, expressed as mg N/kg soil.

For the variables TON (mg kg⁻¹), NH₄ (mg kg⁻¹) (NH₄⁺), soil-extractable Cu (ppm) and Ni concentrations (ppm), a two-way analysis of variance (ANOVA) was performed with the structure: Variable ~ Treatment * Day of Measurement. The residuals of the Ni soil concentration ANOVA were not normal as per the histogram, scatter plot and Shapiro–Wilk test (p < 0.05). As this violated one of the ANOVA assumptions, a Box-Cox test was performed on the model giving the value −1. The reciprocal of the dataset was then used for the statistical analyses. (Note that the transformation did not affect which treatments were significantly different from each other). The transformed residuals Shapiro–Wilk test had a p value of greater than 0.05, indicating normality. Cumulative NO Emissions (kg ha⁻¹) were also analysed using ANOVA with a one-way structure (Variable ~ Treatment). A linear mixed-effect model was performed on the repeated measure variables (¹⁵NH₄⁺, ¹⁵N₂O and ¹⁵NO₃⁻) data using the package lme4 in R. The model was structured as: lmer (Variable ~ Treatment * Days of Measurement + (1 | Experimental Unit)). Tukey’s Honestly Significant Difference test was used to test difference between means for all analyses.

3. Results

3.1. Soil-Extractable Cu, Ni Levels After Application of Metal Salts in Combination with ¹⁵Urea

Overall, the application of CuSO₄ to T2 significantly increased soil levels of available Cu in this treatment (p < 0.001) (Figure 1a) and those levels remained similar in this treatment on the two experimental days analysed (p > 0.05). Maximum levels of 0.24 ± 0.02 ppm were observed in T2 while an average concentration (± standard error, se) of 0.06 ± 0.02 ppm was observed in the rest of treatments on both days analysed.

Figure 1. Concentration of soil-extractable Cu (a) and soil-extractable Ni (b) in soil samples. Bars represent mean values (n = 3) and error bars the standard error of the mean. CK: ¹⁵N-Urea (50% WFPS–75% WFPS); T1: ¹⁵N-Urea + Ni (50% WFPS–75% WFPS); T2: ¹⁵N-Urea + Cu (50% WFPS–75% WFPS); T3: ¹⁵N-Urea + Ni (continuous 75% WFPS).

Soil-extractable Ni (Figure 1b) was significantly higher in the treatments amended with NiSO₄ (T1 and T3) on the days analysed (p < 0.001). No significant differences were observed between the treatments with Ni amendment (T1 vs. T3) (p > 0.005) or between treatments without added Ni (CK, T2) (p > 0.005). Levels of added Ni remained similar on the two dates analysed (p > 0.05). Average values (± se) in T1 and T3 were 1.38 ± 0.11 and 1.77 ± 0.19, while values of 0.70 ± 0.02 and 0.77 ± 0.02 were found in CK and T2, respectively.

3.2. Soil Mineral N Levels and Contribution of the ¹⁵N Urea to Those Levels

The application of NiSO4 (T1 and T3) or CuSO4 (T2) in combination with urea did not significantly change soil NH₄ levels when compared to CK (p > 0.05), but differences were observed between the sampling days (p < 0.05). Urea application rapidly increased NH₄ levels in all treatments, maintaining high values up to 14 days after amendment, and following the same decreasing pattern afterwards (Figure 2a). The maximum soil NH₄ levels, observed from CK, T1, T2 and T3 in the first 14 days of the experiment, were 90.5 ± 9.1, 94.6 ± 11.1, 90.1 ± 5.28 and 71.6 ± 9.9 mg N/kg dry soil, respectively.

Figure 2. Soil NH₄⁺ (a), NH₄⁺ derived from ¹⁵N-urea (b), soil TON (NO₃⁻ + NO₂⁻) (c) and ¹⁵TON (NO₃⁻ + NO₂⁻) derived from ¹⁵N-urea (d) during the experimental period. Data points represent mean values (n = 3) and error bars represent standard error of the mean. Treatments: CK: ¹⁵N-Urea (50% WFPS–75% WFPS); T1: ¹⁵N-Urea + Ni (50% WFPS–75% WFPS); T2: ¹⁵N-Urea + Cu (50% WFPS–75% WFPS); T3: ¹⁵N-Urea + Ni (continuous 75% WFPS).

The soil ¹⁵NH₄ levels analysed showed lower ¹⁵N enrichment of NH₄ in T3 in comparison with the other three treatments on Day 7 and lower than T1 and T2 on Day 14, but the differences were not significant (p > 0.05). With regard to the source of the soil N pool during the experiment, it was observed that a large proportion of the soil N was coming from the amendment on Days 7 and 14 (averages of 72 ± 0.7% and 81 ± 2.7%, respectively), decreasing after these days to average levels as low as 46 ± 1.3% on Day 43 (Figure 2b).

No significant differences were observed in TON levels between treatments (p > 0.05) while differences were observed between sampling dates (p < 0.001). Soil TON concentration gradually increased in all treatments, reaching the maximum levels 29 days after the urea application (Figure 2c). Mean TON levels ranged from 13.1 ± 0.2 (on Day −2) to 80.4 ± 1.1 (on Day 29). The ¹⁵N-TON data showed no differences between treatments (p > 0.05) but there were differences between sampling dates (p < 0.001). The contribution of the ¹⁵N-labelled urea to the soil TON pool gradually increased WITH respect to initial TON levels in all treatments, increasing from average values of 22 ± 0.8% on Day 7 to average values of 57 ± 1.0% on Day 43 (Figure 2d).

3.3. Fluxes of NO and N₂ and Cumulative Emissions

Fluxes of NO (Figure 3) were recorded only during 29 days after the application of the amendments as a problem with the instrument occurred from this day onward, halting the ability to continue monitoring the fluxes for the rest of the experimental period. The magnitude of the measured fluxes from CK, T1, T2 and T3 during the 29 day period were in the ranges 3.6 × 10⁻⁶–7.4 × 10⁻⁵, 3.5 × 10⁻⁶–7.9 × 10⁻⁵, 3.6 × 10⁻⁶–9.8 × 10⁻⁵ and 3.3 × 10⁻⁶–5.2 × 10⁻⁵ kg N ha⁻¹ h⁻¹, respectively. NO emissions showed an initial peak from Day 4 of the incubation in all treatments, reaching maximum fluxes on Day 8 from treatments CK, T1 and T3. Fluxes from T3 were smaller in magnitude when compared to the other three treatments in this initial phase. While NO emissions decreased for treatments CK, T1 and T2, a second peak of NO occurred in T3 from Day 12, reaching its peak at Day 14 followed by a decrease and levelling out by Day 22. In treatments CK, T1 and T2, NO emissions started to increase steadily only a few hours after the addition of water to increase soil moisture levels to 75% WFPS in those treatments. Fluxes of N₂ were not detected during the incubation from any treatment.

Figure 3. Measured NO fluxes after amendment application. Data points represent mean values (n = 3) and error bars represent standard error of the mean. Treatments: CK: ¹⁵N-Urea (50% WFPS-75% WFPS); T1: ¹⁵N-Urea + Ni (50% WFPS-75% WFPS); T2: ¹⁵N-Urea + Cu (50% WFPS-75% WFPS); T3: ¹⁵N-Urea + Ni (continuous 75% WFPS).

The post-hoc statistical analysis showed CK only to be significantly greater than T3 when looking at cumulative NO emissions (p < 0.05) (Figure 4). The ANOVA analysis for cumulative NO emissions produced a p value (p = 0.051) that did not indicate a significant difference between the treatment within the 95% confidence interval. However, this analysis still indicates a high likelihood that the T1 and T2 treatments had a different impact on cumulative NO emissions than T3.

Figure 4. Cumulative NO emissions (kg N ha⁻¹) of the different treatment groups. Boxes represent the datapoints distribution (n = 3), the horizontal line the median value and the whiskers the lower and upper 25% data within each treatment. Treatments: CK: 15N-Urea (50% WFPS-75% WFPS); T1: 15N-Urea + Ni (50% WFPS-75% WFPS); T2: 15N-Urea + Cu (50% WFPS-75% WFPS); T3: 15N-Urea + Ni (continuous 75% WFPS).

3.4. Ratio of N₂O Derived from the Added ¹⁵N-Urea and N₂ Emissions

The ¹⁵N₂O spot samples collected during the experiment showed significant differences in experimental days and the interaction between experimental day and the treatments (Figure 5). The ratio of N₂O derived from the ¹⁵N-labelled urea of all the treatment groups followed a similar increasing pattern after amendment application. Average values ranged from 4.0 ± 0.4% (on Day 0) to 16.5 ± 1.5% (on Day 27). Though not significantly different, the ratios of N₂O derived from the amendment were initially higher in the treatment with the higher level of WFPS (T3—75%). After the soil moisture was increased in CK, T1 and T2, this relationship reversed. While all treatments showed an increase in the ratio of N₂O derived from the amendment, this increase slowed down for treatment T3, which had a lower ratio of N₂O derived from its amendment compared to the other treatments by Day 27.

Figure 5. ¹⁵N atomic ratio in the collected N₂O samples on different experimental days. Data points represent mean values (n = 3) and error bars represent standard error of the mean. Treatments: CK: ¹⁵N-Urea (50% WFPS-75% WFPS); T1: ¹⁵N-Urea + Ni (50% WFPS-75% WFPS); T2: ¹⁵N-Urea + Cu (50% WFPS-75% WFPS); T3: ¹⁵N-Urea + Ni (continuous 75% WFPS).

4. Discussion

4.1. Effect of Application of CuSO₄ on Complete Denitrification

The study aimed to evaluate whether adding CuSO₄ together with urea could affect N cycling. By enhancing soil conditions for Cu-dependent N₂O reductase activity, the aim was to promote total denitrification and, consequently, promote N₂ production. Some studies have reported significant N₂O reductions after CuSO₄ applications. For example, Shen et al. [34] observed 94–95% reductions in N₂O emissions from Andosols and Fluvisols after the application of 130 mM of CuSO₄.

In this incubation study, no N₂ emissions were detected despite the fact that the DENIS system can detect very low levels of N₂ if produced (as low as 0.0004 kg N/ha/h). This absence of N₂ peaks gave us an indication that perhaps no complete denitrification was produced. The treatment that received Cu salt (T2) behaved in the same way as the control treatment (CK), showing no evidence of stimulation of complete denitrification through N₂ production. The daily data provided the %N₂O derived from the ¹⁵N-urea applied from each chamber (Figure 5) as a result of microbial processes that released N₂O, which occurred in all treatments either from soil or from the applied amendment, just not complete denitrification.

In some studies where soil metal applications were at different rates, the focus was on promoting/exacerbating soil conditions to see the effect on N₂O and N₂ production without considering whether those effective rates cause soil and plant toxicity and ultimately an environmental risk. Black et al. [35] reported that enhancing the reduction of N₂O to N₂ requires Cu application rates between 150 and 200 mg/kg, which are above the 20 mg/kg reported to cause plant toxicity. This could explain the similar absence of N₂ production between the treatments despite the application of Cu in T2. In this experiment, the application of CuSO₄ was 8 mg/kg, below the concentration for plant toxicity and much lower than the application by Black et al. [35]. Devaney et al. [36] observed a reduction in N₂O emissions in a sandy loam soil, under similar soil WFPS laboratory conditions (75–80% WFPS) and at low Cu application rates (13.3 mg Cu/kg soil), where daily N₂O flux could reach maximum values of 0.12 kg N/ha from untreated soils while small Cu salt applications produced lower maximum daily values of 0.08 kg N/ha. Sharma et al. [37], during the incubation of riparian, marsh and stream soils with different Cu additions, also observed significant accumulation of N₂O in those soils when no Cu was added in comparison with the N₂O emitted under low (8.3–31.8 mg Cu/kg soil) and high Cu loading (82.6–317.3 mg/kg).

Although N₂O emissions were not measured directly, N₂O was detected from the tracing method. It was also observed that the potential production of this gas could be more related to the optimal soil moisture levels to produce this gas rather than the application of Cu. This can be seen in the increasing trend in enrichment from T3, where no Cu was applied but soil conditions were continuously anaerobic (75% WFPS), and the increase in the ¹⁵N atomic ratio of the N₂O in the rest of treatments, once soil moisture was raised to anaerobic conditions.

In addition to the low Cu application rate, it is possible that a large proportion of the Cu applied with CuSO₄ was retained by the large fraction of clay present in the soil used for the experiment (Cambisol). The capacity of adsorption of Cu increases with the higher levels of clay minerals [26,38]. Besnard et al. [39] found larger concentrations of Cu in clay-sized fractions of soils, particularly clay size fractions <2 µm. Thus, it is possible that, by bonding with clays, Cu became less mobile and less available as a nutrient, as reflected in the low extractable Cu levels observed in the soil (Figure 1a). Under conditions of environmentally safe CuSO₄ application rates and low Cu availability in the Cambisol studied, it is possible that the expected effect of the added Cu to increase N₂ emissions through increased activity of N₂O reductase was counteracted by the reduced availability of Cu after its application through binding processes. Further incubations need to be conducted to clarify this.

4.2. Effect of Application of NiSO₄ on Nitrification

Urease is a Ni-dependent enzyme that catalyses urea hydrolysis into NH₃, which is transformed to NH₄⁺ when in equilibrium with water, which can then be nitrified. Some studies have reported that Ni additions to soils can stimulate nitrification. deCatanzaro et al. [40] observed this effect on forest soils after applications of 100 mg/kg soil. In our study, no clear increase in NO production due to NiSO₄ was observed, at least during the period of 29 days monitored by the NO analyser. NO emissions from T1 (NiSO₄) and CK (no NiSO₄) were similar at soil moisture levels (50% WFPS) where nitrification occurred (first 14 days). This was reflected by the similar increase in soil NH₄ levels and the gradual transformation to soil NO₃ in both treatments (Figure 2). The absence of differences between T1 and CK can also be attributed, as for CuSO₄, to the low NiSO₄ application rates applied based on the Government’s recommendations and to the clay content of the soil used for the incubation [41,42]. While several studies have investigated the effects of heavy metals on the soil N-cycle, specific studies investigating the resulting NO emissions from metal applications to soils were not found in the literature, reflecting the need for more studies to investigate the effect of this metal in different type of soils and experimental set-ups.

4.3. Effect of Moisture Conditions and Metal Application on NO and Potential N₂O Emissions

The Ni treatment at a constant high WFPS level (T3) showed the lowest cumulative NO emissions. A comparison of the treatments with added NiSO₄ (T3 and T1) indicates that the differences in NO production between treatments (despite being not significant at exactly 95% confidence) are highly likely to be driven by the moisture conditions [18,19] rather than metal application, at least during the period monitored. It is important to remember that NO, produced by nitrification and denitrification processes, is not always emitted as it can be rapidly consumed in the soil, mainly during the denitrification process where relative consumption is higher [43]. Since this occurs under anaerobic conditions, it is plausible to believe that the more anaerobic conditions in T3 led to greater consumption of NO than T1. Higher NO emissions could suggest higher potential for N₂O emissions, which could have started to be produced from Day 13, as NO emissions decreased [44]. The significantly higher contribution of the applied urea to the ¹⁵N ratio of the N₂O samples collected in CK, T1 and T2 in comparison to T3 from Day 14 also coincided with the increase in soil moisture in those treatments (Figure 3). This seemed to have a more significant impact on the transformation of the applied amendment to N₂O than the application of heavy metals to soils. At this stage, soil mineral N data also indicate increasing soil NO₃ levels, which, under the more anaerobic soil conditions, support the theory of higher N₂O emissions due to denitrification at this stage. Equally, if the dry soil is rewetted, any N₂O produced and stored in the soil could then be released. Thus, the change in soil moisture could affect the timing of the N₂O peaks but may not have significantly reduced the overall N₂O emission, when the soil was highly moist for a certain period. However, this inference still requires evidence from further studies looking at N₂O emission data. It is very important to note that the NO analyser did not acquire data after Day 29, where soil moisture levels were still anaerobic, so any potential effect of Ni application on NO produced from this soil after this day was missed, and therefore, it was not possible to detect or evaluate.

The addition of Ni and Cu (represented by T1 and T2, respectively) did not significantly affect the source of the ¹⁵N atomic ratio of the N₂O spot samples compared to the CK treatment, at any of the sampling times, suggesting only a minor impact on the potential N₂O emissions. Chiaravalloti et al. [45] also observed that the application of metals such as Cu can significantly decrease N₂O emissions, but this effect depends not only on the rate of Cu applied but alos on the soil pH, being significant under a high soil pH but not in acidic soils with low pH. Only microorganisms that have nosZ genes encoding N₂O reductase can conduct a complete reduction of N₂O to N₂. At low soil pH, the synthesis, activity and functionality of N₂O reductase are inhibited, favouring N₂O emissions [46]. Shaaban et al. [47] observed that reductions in N₂O emissions from acid soils are possible under adequate Cu levels, increased pH and abundant nosZ genes in those soils. In this sense, the acidic nature of the soils used for the incubation in combination with the use of the UK government’s Cu recommended rates (low) could have been factors explaining the lack of an effect on N₂ emissions. deCatanzaro et al. [40] observed that high Ni applications (500 mg/kg) stimulated N mineralisation, and these rates were much higher than those used in our study. However, mineralisation processes are likely to have occurred during the experiment in all treatments. Initial soil mineral N contents on Day −2 were significantly lower than those of the samples after the addition of the ¹⁵N-urea. When comparing the levels of NH₄⁺ with the % soil ¹⁵NH₄⁺ from amendment (and ¹⁵NO₃⁻) (Figure 2a–d), differences between 20 and 30 g N/kg of soil were observed, denoting a possible contribution from mineralisation to increased available soil N. Mineralisation processes were likely to happen in this soil since the experiment was conducted in sieved (2 mm) and air-dried soil to reach a WFPS level of 30%. Under these conditions, the organic matter (initially protected in the soil aggregates before sieving) was more physically exposed and unprotected. Thus, the rapid soil rewetting to establish experimental moisture conditions could have triggered mineralisation [48] by the microbial community in all treatments and increased the soil-available N pool.

4.4. Recommendations for Future Research

The availability of metals in the soil can regulate or inhibit microbial processes by causing changes in the microbial population, ultimately altering N-cycling. These changes can be observed after short-term manipulation of the metal availability [24]. In this incubation, the soil type use, in combination with the selected UK Government’s recommendation rates (realistic rates for agronomic use), may have limited the ability to produce an enzymatic effect that translated to evident differences between treatments due to the low metal availability for microbes. In addition to the low metal application rates applied to this soil, the high clay content may have further reduced the amount of metal availability for microbial activity by adsorption to this fraction. In this sense, future experimental designs that include different application rates and different soil types should be conducted to untangle the potential dual effect of these two factors (rate, soil type) seen in this incubation. Black et al. [49] reported that some denitrifying bacteria are sensitive to historical concentrations of Cu in soils. This incubation only covered a period of 43 days after a single application event. Hence, the potential effects of the long-term exposure to these metal rates in this soil on microbial activity and N-cycling should also be addressed in future incubations. In relation to moisture conditions, the current incubation included treatments coupled with soil moisture conditions and metal applications. For Cu, it was not possible to fit a treatment with continuous anaerobic conditions (%75 WFPS) so future designs should aim to decouple moisture conditions and metal effects from both Cu and Ni, with full factorial structure and control.

We acknowledge the lack of continuous N₂O fluxes during this incubation and the importance of these being monitored in future incubations to observe the clear effect of the metals on the N dynamics that lead to its production or reduction to N₂. In relation to this, and as mentioned before, the influence of Cu availability on N₂O consumption/reduction (even if N₂O data were available) in this soil would have been better evaluated with experiments conducted at microbiological and enzymatic levels. For Ni, urease activity, urea hydrolysis rates and changes in NH₄⁺ specific to Ni applications were not measured in this study so future studies should include these to obtain a better understanding of processes that lead to NO emissions.

Shen et al. [34] observed that only 54% of the strains of denitrifiers investigated from an Andosol and a Fluvisol were able to reduce the N₂O to N₂ after application of Cu to pure culture, showing the complexity of the soil processes involved. Therefore, similar experiments should be conducted with this Cambisol to elucidate the potential of Cu or Ni applications to reduce N losses at the agronomic levels recommended by the UK Government.

5. Conclusions

Following the UK Government’s recommended soil application rates of Cu and Ni salts is crucial for providing nutrients for biological plant processes without causing soil toxicity from a excess application. However, the effect of soil N emissions needed to be examined. The data acquired with this soil incubation experiment showed that the application of these metals at the recommended rates do not seem to stimulate N processes that could affect NO emissions or promote losses of N through N₂ emissions in Cambisols. The effect of these processes could have been counteracted by the reduced availability of the metals after their application through binding processes with the clay fraction as well as the acidic nature of the soil. However, results showed that there was a high likelihood that the resulting NO emissions were related to the different soil moisture conditions between treatments. Further research into the effect of metal applications on gaseous N emissions from contrasting soil types, application rates, repeated application events and soil moisture conditions is required. Future studies in this type of soil should be conducted in parallel with microbial and enzymatic experiments to elucidate the specific effect of these metal application rates on the specific microbial population and the resulting N gases.