Influence of Sediment, Plants, and Microorganisms on Nitrogen Removal in Farmland Drainage Ditches

Abstract

The removal of nitrogen from water is a consequence of the synergistic action of plant uptake, sediment sorption, and microbial decomposition. However, there is a lack of long-term experimental studies on the effects of each component in the process of nitrogen removal. In this study, we investigated the effect of sediment, plants, and microorganisms on nitrogen removal by setting up three systems: water–sterilized sediment, water–sediment, and water–sediment–plant. The nitrogen removal effect followed the following rank order of effectiveness: the “water–sediment–plant” system > the “water–sediment” system > the “water–sterilized sediment” system. The ditch sediment had a strong enrichment effect for nitrogen. In addition, the migration rate of nitrogen in the sediment with different depths was different. The ammonia-nitrogen migration rate in the sediment showed an increasing trend with time and depth. The nitrate-nitrogen migration process in the sediment showed a trend of enrichment toward the middle layer (15.0–25.0 cm). Aquatic plants and microorganisms can promote the removal of nitrogen in water, with the average purification rates of 13.92% and 19.92%, respectively.

1. Introduction

Water pollution is a very important issue [1]. With the improvement in public awareness and the strengthening in supervision, point source pollution is gradually being brought under comprehensive control, and an increasing number of people are beginning to pay attention to agricultural non-point source pollution, especially the pollution caused by the heavy use of chemical fertilizers and pesticides [2]. The process of agricultural non-point source pollution, starting from the formation of farmlands to the output of water bodies outside the farmland, can be partitioned into two phases: first, the production of pollutants—such as N, P, and others—in fields with surface runoff, subsurface seepage, and other processes into a final drainage ditch; second, the output of pollutants to external water bodies through the branch and dry-level farmland drainage ditches. The application of ecological engineering techniques to design or modify agricultural drainage ditches can effectively control agricultural non-point source pollution [3,4]. Furthermore, due to the low construction cost of eco-drainage ditches, they are more suitable for promotion and application in rural areas [5,6,7].

Nitrogen in agricultural drainage ditches exists in both organic and inorganic forms; however, the mechanism of nitrogen removal is complex because of the various subdivisions of inorganic nitrogen and the conversion process of the redox reactions between different forms [8]. The transformation of nitrogen in ditches occurs mainly through two balance processes: the balance of deposition and mineralization with organic nitrogen [9] and the balance of sediment sorption, plant uptake, microbial absorption, nitrification, and denitrification in the form of ammonia and nitrate-nitrogen [10,11,12,13]. The removal of nitrogen from water is the result of the synergistic action of plant uptake, sediment adsorption, and microbial decomposition. Plants can directly absorb nitrate-nitrogen (NO₃⁻-N) and ammonia-nitrogen (NH₄⁺-N) from water. In addition, plant roots can provide an alternating aerobic and anaerobic environment for nitrification and denitrification, which promotes the conversion and removal of nitrogen and its assimilation into plants. Ditch sediments can directly remove NO₃⁻-N and NH₄⁺-N from water bodies through adsorption, while various nitrifying and denitrifying bacteria grow in ditch sediments, which can effectively remove the nitrogen from water via the synergistic effect of physical adsorption and biochemical action [14].

The current understanding of the mechanism of nitrogen transport and transformation by plants is mainly achieved through in situ monitoring experiments, such as the screening and planting of aquatic plants [15,16]. Studies on microbial purification mechanisms are mostly conducted through nitrification and denitrification mechanisms, as well as the addition of microbial strains. Zhao et al. [17] showed that ammonia-oxidizing microorganisms, particularly ammonia-oxidizing archaea (AOA) are suitable for the treatment of nitrogen-containing wastewater. Studies on the purification mechanism of sediments have been conducted through adsorption and desorption experiments, as well as in situ monitoring [18]. In addition, Chen et al. [19] found that the combination of submerged plants and microorganisms had a positive effect on total nitrogen (TN) and total phosphorus (TP) removal in marginally polluted water, with removal rates of 70% and 97.3%, respectively. The planting of reeds considerably reduces TN, total dissolved nitrogen, and NH₄⁺-N in the sediment, as well as promotes the mineralization, nitrification, and denitrification of organic nitrogen in the sediment [20]. In summary, there are many studies on the mechanisms of nitrogen purification by sediment, plants, and microorganisms in drainage ditches, but they are limited by experimental conditions and methods, and only few long-term studies are available.

There are many experimental studies and model simulations on the purification effects, processes, and degradation mechanisms of pollutants in agricultural drains [21,22,23]; for example, Kroger [24,25] found that a 400 m long and 7 m wide agricultural ditch could retain 57% of soluble inorganic nitrogen. Fan et al. [26] developed a nitrogen balance model by analyzing the changes in TN, NH₄⁺-N, and NO₃⁻-N content in the whole system, providing a new idea for enhanced nitrogen removal from drainage ditch systems. Zhan et al. [27] used the HYDRUS-1D model to simulate the removal of NH₄⁺-N from soil based on an indoor soil-column leaching experiment. Huang et al. [8] used the SWAT model and a modified SWAT-N₂O coupler to investigate the effects of climatic factors on the transport and transformation of non-point source nitrogen and N₂O emissions in an agricultural drain ditch. However, most studies were conducted to evaluate the overall purification effects of drains as a system or model simulation.

Studies in this area have deepened the understanding of the processes and effects of nitrogen purification in agricultural drainage ditches. However, due to the challenge of distinguishing the individual contributions of plants, sediments, and microorganisms in the removal of nitrogen within drainage ditch systems, their specific roles in the nitrogen removal process have not been adequately investigated [28,29], and there is a lack of long-term experimental studies on the effects of each component on nitrogen removal. Therefore, to address this research gap, this study conducted a meso-scale experimental study of microorganisms, sediments, and plants so as to understand their effect on the nitrogen migration and transformation process for about seven months by setting up three systems: the “water–sterilized sediment”, “water–sediment”, and “water–sediment–plant” columns. We hypothesized that (1) the purification effects of plants, sediments, and microorganisms on nitrogen in water can be superimposed, and (2) that ditch sediment has a strong enrichment effect for nitrogen. In addition, the migration rate of nitrogen in sediments with different depths was different. This study has important research significance for understanding the mechanism and influencing factors of nitrogen removal in farmland drainage ditches, and it can provide a theoretical reference and experimental basis for alleviating agricultural non-point source pollution.

2. Materials and Methods

2.1. Experimental Design

The experimental equipment was set up at the Water Conservation and Agroecology Experimental Base of the Yellow River Institute of Water Resources Science (35°18′ N, 113°53′ E). The area has a temperate continental monsoon climate, with an altitude of 73.2 m, an average annual temperature of 14.1 °C, a frost-free period of 210 d, 2398.8 h of sunshine, and an average annual precipitation of 582.0 mm (with June to October accounting for 70.0–80.0% of the annual rainfall).

Three Plexiglas columns (with a 60 cm inner diameter and 100 cm height) were used for the experiment, with the lower end closed and the upper end open. A 90 cm ranged transparent ruler was pasted on the glass column. The depths of the sediment and overlying water in the column were 40 and 50 cm, respectively. Sediments were collected from the experimental base. The water used for the experiment came from the groundwater in the well at the base, and the depth of the overlying water varied between 10 and 50 cm during the experiment (water was replenished when the water level dropped to 10 cm due to evaporation). The nitrogen index of the injection groundwater during the experimental period is shown in Table 1. The particle composition of the sediments in the study was 7.74% (<2.00 μm), 60.59% (2.00–50.00 μm), and 31.67% (2.00–50.00 μm). The initial nitrogen index of the sediment after one week of introducing the water is shown in Table 2. The WS column represents a “water–sediment” system, and the WSP column a “water–sediment–plant” system with one well-developed reed that was planted on the substrate. In order to study the purification effect of soil microorganisms on nitrogen, it is necessary to set a blank control and to sterilize the sediment so as to compare the purification effect of the sterilized sediment and the sediment on nitrogen. The WSS column represents a “water–sterilized sediment” system, where the sediment is autoclaved before filling (Figure 1). At the same time, to prevent the influence of external weather (rain, snow, and other precipitation), as well as the entry of mosquitoes and external nitrogen sources to the water body, the columns were placed in bird-proof cages and covered with rainproof covers during precipitation.

2.2. Experimental and Analytical Methods

This experiment was conducted from 7 April 2016 to 21 November 2016. Water samples (30 mL) were collected every two days in the early stage of the experiment and once a week in the later stage, with an additional measurement after each recharge. This was achieved using an injection syringe to aspirate the overlying water at half depth while also measuring the pH and temperature of the overlying water. Sediment samples were taken once a month at 0.0–10.0 cm, 10.0–20.0 cm, 20.0–30.0 cm, and 30.0–40.0 cm. Before measurement, the water samples were shaken in the sampling bottle, and then filtered in a conical bottle through the phosphorus-free filter paper. The soil samples were ground through a sieve, where each soil sample weighed 3 g, and were placed in a 100 mL conical flask. Then, 60.0 mL (at 2.0 mol/L) of KCl solution (at a 1:5 soil:KCl solution ratio) was added. The samples were then sealed with a sealing film and oscillated on a constant temperature oscillator for 1 h. Next, the samples were removed and filtered, and the filtrate for then taken for measurement. The NH₄-N and NO₃⁻-N contents in the water and sediment were directly measured by a Smartchem 140 automatic chemical analyzer (Alliance Corp., Paris, France). A HYDROLAB HL7 multi-parameter water quality analyzer (Hach, Loveland, CO, USA) was used to measure water parameters (temperature, pH, electrical conductivity, etc.).

As shown in Figure 1, when the other experimental conditions were the same, the difference in nitrogen concentrations in column WSP and column WS can be attributed to role of plants in the nitrogen migration and transformation process. As the NH₄⁺-N in water was assimilated by plants, adsorbed by sediment, and degraded by microorganisms (in addition to being easily converted into ammonia gas and volatilized under alkaline conditions), the NO₃⁻-N in the WS and WSP water columns was chosen as the study object. Similarly, when the other experimental conditions were the same, the difference in nitrogen concentration in the WSS and WS columns could be attributed to the role of microorganisms in the process of nitrogen transport and transformation. As the NO₃⁻-N was both a generator of nitrification and a reactant of denitrification, the change in NO₃⁻-N concentration could better reflect the role of microorganisms. To quantify the process of nitrogen migration and transformation, the plant purification rate W_p (WSP column vs. WS column) and the microbial purification rate W_m (WS column vs. WSS column) were calculated separately. The calculation equations are shown in Equations (1) and (2):

(1)$W_P=(C_{ws}-C_{wsp})/C_{ws}\times 100\%$

(2)$W_m=(C_{ws}-C_{wss})/C_{ws}\times 100\%$

where W_p is the plant purification rate (%), W_m is the microbial purification rate (%), and C_ws, C_wsp, and C_wss are the nitrogen concentrations in the water column in mg/L.

In addition, to quantify and analyze the degree of nitrogen transport and transformation between the sediments at different depths, the sediment migration rate M_a–b from depth a to depth b was calculated by Equation (3):

M_5–15 = (C_10–20 − C_0–10)/C_10–20 × 100%(3)

where M_5–15 is the sediment migration rate from a depth of 5.0 cm to 15.0 cm (%), and C_0–10 and C_10–20 are the nitrogen contents (g/kg) in the sediment at 0–10.0 cm and 10.0–20.0 cm, respectively.

The particle compositions of the sediment were 9.58% <2.00 μm, 60.48% 2.00–50.00 μm, and 29.94% >50.00 μm.

2.3. Statistical Analyses

SPSS 24.0 (IBM, New York, NY, USA) and Origin 2021 9.8.0.200 (Origin Lab, Northampton, MA, USA) software were used for data processing, graphing, and tabulation. Each data point was summarized by calculating the average value and standard deviation (S.D). A one-way analysis of variance (ANOVA) was conducted to compare the effects of the “water–sediment”, “water–sediment–plant”, and “water–sterilized sediment” systems on nitrogen concentration within the same time period. A significance level of p = 0.05 was set. And Origin 2021 was used for the linear regression analysis of the relevant data.

3. Results

3.1. Variation of pH, Electrical Conductivity, and Temperature in the Overlying Water

The variations in pH during the experiment were as follows: the WS column at 7.03–9.64 with a mean value of 8.98; the WSP column at 7.96–9.75 with a mean value of 9.00; and the WSS column at 7.94–9.82 with a mean value of 9.14. The variations in water temperature during the experiment are shown in Figure 2b. The range and trend of temperature changes in the WS–WSS columns were essentially the same during the experiment, showing an initial rise (7 April–21 June), followed by a fluctuation (21 June–29 August), and then a decline (29 August–30 November). The variation in temperature was in the range of 8.02–31.8 °C, with an average of 21.8 °C. The highest and lowest temperatures were recorded on 29 July and 30 November, respectively. The variation in electrical conductivity is shown in Figure 2c: the WS column at 1661.0–3820.0 mS/m with a mean value of 2501.8 mS/m; the WSP column at 1564.0–3750.0 mS/m with a mean value of 2397.2 mS/m; and the WSS column at 1620.0–3450.0 mS/m with a mean value of 2357.5 mS/m. During the experiment, the electrical conductivity showed an increasing trend (7 April–18 October), gradually reaching a peak, and then decreasing (18 October–16 December).

3.2. Variation in Nitrogen in the Overlying Water

The purification process of the NO₃⁻-N in the WS, WSP, and WSS columns is shown in Figure 3. After groundwater was injected into the three experimental columns on 7 April, the concentration of NO₃⁻-N in the WS to WSS columns rose rapidly to reach the first peak of 18.60, 23.09, and 19.73 mg/L on 14 April, indicating the release of NO₃⁻-N from the sediment to the overlying water through the interstitial water. The highest concentration of NO₃⁻-N in the WSP column may be because, during the reed transplanting process, the reed root sediment contained more organic matter, which was released into the overlying water through the interstitial water. The content of NO₃⁻-N in the WS, WSP, and WSS columns showed a similar trend, with the concentration increasing after groundwater injection on 7 April followed by a decline. A similar trend was shown on 19 April, 5 May, 26 May, 12 June, 21 June, and 3 November following the recharge of groundwater. Similarly, disturbances caused by the collection of sediment samples on 3 June, 22 July, 15 August, 14 September, and 18 October also resulted in a transient increase and decrease in the concentration of nitrate nitrogen in the water.

The concentration of NO₃⁻-N in the WS–WSS columns decreased from 14 April to 1 July, and followed the following rank order: WSP column (“water–sediment–plant” system) < WS column (“water–sediment” system) < WSS column (“water–sterilized sediment” system). The nitrate nitrogen content in the WS, WSP, and WSS systems all showed a decreasing trend at the initial stage, and the rate and range of the nitrate nitrogen content decrease in the WSP system was the largest (Figure 3).

From 1 July to 13 October, the NO₃⁻-N concentrations fluctuated at a low level with a range of ≤2.79 mg/L and a mean concentration of 1.45 mg/L. The two peaks of the NO₃⁻-N concentration in the water column occurred on 18 October and 3 November, respectively. The peak in the water column occurred on 18 October, which may be attributed to the leakage of the soil from the sampler during soil extraction, which increased the NO₃⁻-N content in all three water columns. On 3 November, the increase in NO₃⁻-N concentration may be attributed to the weakened plant absorption capacity and the weakened microbial activity.

Figure 4 shows that the variation in the NH₄⁺-N in the WS–WSS columns was similar to that in the NO₃⁻-N. Both the water column recharge and sediment sampling increased the concentration of ammonia in the water column, with a fluctuating upward trend from 7 April to 1 July; however, it remained stable with small changes from 1 July to 13 October, but then a fluctuating upward trend occurred from 18 October to 21 November. The average NH₄⁺-N concentration in the columns during the entire experimental period followed the following rank order: WSP column (0.32 mg/L) < WS column (0.44 mg/L) < WSS column (0.47 mg/L). A one-way ANOVA analysis showed that there was no significant difference in the variation in NH₄⁺-N concentration in the water columns from the WS–WSS systems, while there was a significant difference in the variation in NO₃⁻-N in the columns, as shown in Table 3. This is because the presence of the water column above the sediment resulted in an anaerobic environment that facilitated denitrification, which is the main reaction process by which soil microorganisms reduce NO₃⁻-N to nitrogen gas or nitrogen oxide. The reaction substrate is NO₃⁻-N, which disrupts the NO₃⁻-N balance between water and soil, as well as causes considerable differences in the NO₃⁻-N processes in the water column [30].

3.3. Variation in Nitrogen in the Sediment

To investigate specifically the sedimentation and absorption of the nitrogen by the ditch sediment, excluding the role of microorganisms and plants, the WSS column (water–sterilized sediment) was chosen for study. The changes in the NH₄⁺-N content and the migration rate in the sediment at different depths are shown in Figure 5. The NH₄⁺-N content in the sediment at different depths showed an increasing trend during the operation period. In May and June, as the temperature increased, the adsorption of the sediment was enhanced and the NH₄⁺-N content increased rapidly, indicating the characteristics of a “rapid adsorption” of NH₄⁺-N in the sediment and the maintenance of a “Slight rise” state from August to October. With increasing depth, the NH₄⁺-N content showed an increasing trend, with an average of 9.0 × 10⁻³, 1.0 × 10⁻², 1.1 × 10⁻², and 1.3 × 10⁻² g/kg at 0.0–10.0 cm, 10.0–20.0 cm, 20.0–30.0 cm, and 30.0–40.0 cm, respectively. There was a significant difference in the migration rate of the NH₄⁺-N at different depths (p = 0.00 < 0.05), indicating a trend of enrichment in the deeper layers. This observation was consistent with the mean values of the migration rate of NH₄⁺-N, which were 16.20%, 11.54%, and 10.97%. Further, the migration rate of NH₄⁺-N in the sediment at different depths showed a decreasing trend in the later stage of the experiment, indicating that the migration process gradually weakened.

The variation, as the depth increased, in the NO₃⁻-N content in the sediment at different depths (Figure 6) deviated from the trend observed for the NH₄⁺-N content, with the highest NO₃⁻-N content being represented at a 20.0–30.0 cm depth. The migration rate of the NO₃⁻-N in the substrate at different depths showed the following rank order: 15.0–25.0 cm > 5.0–15.0 cm > 25.0–35.0 cm. The migration rate at a 25.0–35.0 cm depth was negative, indicating that the NO₃⁻-N in the “water–sterilized sediment” system tended to be enriched in the middle substrate, which is consistent with the highest NO₃⁻-N content in the substrate at a 20.0–30.0 cm depth.

4. Discussion

4.1. Effect of Temperature and pH on the Nitrogen Content in Water and Sediment

Investigations have demonstrated that microbial activity and the rate of organic matter degradation are influenced by pH levels, with both excessively high and low values exhibiting an impact. Furthermore, different microorganisms display varying degrees of adaptability to different pH conditions. For nitrifying bacteria, the optimal pH range for nitrification reactions is typically observed between 7.0 and 8.0, while denitrifying bacteria (which play a key role in the denitrification processes), exhibit optimal performance at pH levels ranging from 7.0 to 7.5 [31]. In this study, the pH values of the water samples analyzed fell within this specified range, with all measurements indicating a neutral to weakly alkaline nature. Such conditions are highly conducive to the efficient removal of nitrogen from water.

From April to June, the nitrogen removal rate increased rapidly with increases in temperature. This was because appropriate high temperatures facilitate the microbial nitrification and denitrification reactions for removing nitrogen from the ditches [32]. From June to August, the purification rate of plants and microorganisms showed an oscillating trend with increases in temperature, which was due to the sensitivity of plants and microbial growth to external factors [32,33]. At high temperatures, the growth of plants, as well as the nitrification and denitrification of microorganisms were inhibited to a certain extent, thereby decreasing the nitrogen removal rate; moreover, the microbial population may reach a peak at this time due to the nutrients in the substrate that may not be sufficient to sustain the growth of the population, leading to a dynamic equilibrium [34,35].

In October, the purification rate of the plants and substrate was close to or equal to zero, and after October, the overall purification rate of the plants and microorganisms was less than zero. The desorption of the sediment, decomposition of the roots of aquatic plants, and the degradation of branches and leaves by microorganisms lead to the release of nitrogen into the water, causing secondary pollution [36]. Reduction in temperature had a significant impact on nitrogen removal efficiency. Tan et al. [37] also reported that temperature has a significant impact on soil nitrification; the higher the soil temperature within a certain range, the faster the soil nitrification rate—reaching a maximum at 30 °C (after which soil nitrification is inhibited). However, the optimum temperature for nitrate and nitrogen removal in this experiment was approximately 22 °C, which was lower than that for soil nitrification, probably because of the differences in the optimum temperature for soil nitrification in different properties and regions [38,39], or due to the controlled conditions in this experiment.

4.2. Effect of Plants on Nitrogen Migration and Transformation

The purification rate of NO₃⁻-N by plants showed an overall trend of first increasing, then fluctuating around zero, and finally becoming less than zero (Figure 7). This corresponded to the growth process of the reeds, which began to appear in April. This was then followed by rapid stem growth, and then finally entering into the aging stage. Throughout the experiment period, the average purification rate of the plants was 13.92%, indicating that the aquatic plants overall promoted the removal of nitrogen in the water. However, the influence of plants on nitrogen transport and transformation processes does not solely depend on the plants themselves, but also on the soil microbial community. This is because plants can alter the structural and functional properties of microbes, as well as promote their growth when grown in a new environment [40]. For example, during the peak period of plant growth, the plant N uptake rate is negatively correlated with the microbial biomass N content [41]. This may be but one reason for the fluctuation of the nitrogen migration rate nearing zero from June and October.

4.3. Effect of Microorganisms on Nitrogen Migration and Transformation

The main mechanism of pollutant degradation in ditches is the decomposition and transformation of nutrients by microorganisms [42]. As shown in Figure 8, the purification rate of microorganisms had a similar trend to that of plants with an increasing trend from April to July, which was related to the biochemical effect of microorganisms that were exacerbated by the increase in temperature. After October, the purification rate of microorganisms was less than or equal to zero, showing the same pattern as the purification rate of the plants. This was related to the decrease in temperature, the weakening of microbial activity, and the desorption of the substrate. The purification rate curve of microorganisms showed a sharp fluctuation in July and September because the microorganisms introduced in the process of groundwater replenishing gradually multiplied, resulting in a gradual increase in microorganisms in the WSS column and a reduction in the difference with the WS column. Simultaneously, the biochemical action of the microorganisms was also sensitive to the impact of the external factors [43,44], and this was inhibited under inappropriate conditions [45]. Throughout the experiment period, the average purification rate of the microorganisms was 19.92%, indicating that the presence of microorganisms had a facilitating effect on the purification process of nitrogen in water.

Ammonia nitrogen gradually deposits during the adsorption of ditch sediment [46,47], thus migrating toward deeper layers with time. In addition, it is difficult to avoid the introduction of microorganisms during the groundwater injection process, and different depths of sediment have varying denitrification potentials [48]. Nitrate nitrogen gradually accumulates in the bottom layer, and there may be strong denitrification at a depth of 20–30 cm in the sediment. This results in the consumption of nitrate nitrogen [49], and the indication of a tendency to accumulate in the middle layer. Due to limitations in experimental conditions, this study was unable to obtain the distribution of nitrate, oxygen, and microorganisms in the pore water of the sediment, and thus it cannot specifically discuss the distribution of the vertical denitrification potential of the sediment in this experiment. We will take this into consideration in future research.

5. Conclusions

In this paper, we analyzed the migration and transformation of nitrogen in drainage ditches by setting up three systems—namely “water–sediment”, “water–sediment–plant”, and “water–sterilized sediment”—as well as investigated the role of sediment, plants, and microorganisms in the process of nitrogen purification. This experiment generated the following conclusions.

• (1). In general, the rank order of nitrogen removal in the systems was as follows: the “water–sediment–plant” system > the “water–sediment” system > the “water–sterilized sediment” system.

• (2). The ditch sediment had a strong enrichment effect for nitrogen. The NH₄⁺-N migration rate in the sediment showed an increasing trend with time and depth. The NO₃⁻-N migration process in the sediment showed a trend of enrichment toward the middle layer (15.0–25.0 cm).

• (3). Compared with the WSS system, both the WSP and WS systems showed a better nitrogen purification effect. Aquatic plants and microorganisms can promote the purification of nitrogen in water, with average purification rates of 13.92% and 19.92%, respectively.

Author Contributions

Conceptualization, C.G. and Q.L.; methodology, Q.Z. (Qisen Zhang) and C.G.; investigation, Y.H. and Q.Z. (Qiang Zhao); resources, J.W.; data curation, Q.Z. (Qisen Zhang) and J.W.; writing—original draft preparation, C.G. and J.W.; writing—review and editing, Q.L. and J.W.; funding acquisition, J.W. and C.G. All authors have read and agreed to the published version of the manuscript.

Data Availability Statement

The datasets generated during and/or analyzed during the current study are not publicly available due to the confidential nature of the data but are available from the corresponding authors upon reasonable request.

Conflicts of Interest

The authors declare no conflict of interest.

Footnotes

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Figures and Tables

Figure 1. Schematic diagram of the experimental equipment (unit: cm).

Figure 2. Variation in water quality parameters in the WS–WSS columns: (a) temperature; (b) electrical conductivity.

Figure 3. Variation in the nitrate nitrogen content over time in the three systems (WS, WSP, and WSS).

Figure 4. Variation in ammonium nitrogen content over time in the three systems (WS, WSP, and WSS).

Figure 5. Variation in the NH4+-N content and migration rate in sediments at different depths in the WSS column: (a) NH4+-N content; (b) migration rate of the NH4+-N content.

Figure 6. Variation in the NO3−-N content and migration rate in sediments at different depths in the WSS column: (a) NO3−-N content; (b) migration rate of the NO3−-N content.

Figure 7. Variations in the plant purification rate.

Figure 8. Variations in the microbial purification rate.

Table 1

The nitrogen concentration of the injection groundwater during the experimental period.

Nitrogen Form	Time
	7 April 2016	27 April 2016	5 May 2016	6 August 2016	3 November 2016
NO3−-N (mg/L)	1.7 × 10	1.6 × 10	1.9 × 10	1.5 × 10	1.3 × 10
NH4+-N (mg/L)	2.1 × 10−1	2.4 × 10−1	1.3 × 10−1	3.6 × 10−1	4.2 × 10−1
SUM (mg/L)	1.7 × 10	1.7 × 10	1.9 × 10	1.5 × 10	1.4 × 10

Table 2

Initial nitrogen concentration in the sediment.

Nitrogen Form	WS(Water–Sediment)	WSP(Water–Sediment–Plant)	WSS(Water–Sterilized Sediment)
NO3−-N (mg/L)	2.7 × 10−2	2.9 × 10−2	4.0 × 10−2
NH4+-N (mg/L)	8.0 × 10−3	8.0 × 10−3	8.0 × 10−3
SUM (mg/L)	3.5 × 10−2	3.7 × 10−2	4.8 × 10−2

Table 3

One-way ANOVA of the variation in NH₄⁺-N concentration in the WS–WSS columns (F: variance ratio; P: significance level).

	Time	F	p
Nitrate-nitrogen	7 April 2016–1 July 2016	5.4 × 100	6.0 × 10−3
	7 April 2016–21 November 2016	3.1 × 100	4.7 × 10−2
Ammonia-nitrogen	7 April 2016–1 July 2016	9.0 × 10−2	9.2 × 10−1
	7 April 2016–21 November 2016	7.7 × 10−1	4.6 × 10−1

References

1. Maguire, T.J.; Spencer, C.; Grgicak-Mannion, A.; Drouillard, K.; Mayer, B.; Mundle, S.O.C. Distinguishing point and non-point sources of dissolved nutrients, metals, and legacy contaminants in the Detroit River. Sci. Total Environ.; 2019; 681, pp. 1-8. [DOI: https://dx.doi.org/10.1016/j.scitotenv.2019.04.311] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/31102810]

2. Rissman, A.R.; Carpenter, S.R. Progress on Nonpoint Pollution: Barriers & Opportunities. Daedalus; 2015; 144, pp. 35-47.

3. Coban, O.; Kuschk, P.; Kappelmeyer, U.; Spott, O.; Martienssen, M.; Jetten, M.S.M.; Knoeller, K. Nitrogen transforming community in a horizontal subsurface-flow constructed wetland. Water Res.; 2015; 74, pp. 203-212. [DOI: https://dx.doi.org/10.1016/j.watres.2015.02.018]

4. Liu, F.; Xiao, R.L.; Wang, Y.; Li, Y.; Zhang, S.L.; Luo, Q.; Wu, J.S. Effect of a novel constructed drainage ditch on the phosphorus sorption capacity of ditch soils in an agricultural headwater catchment in subtropical central China. Ecol. Eng.; 2013; 58, pp. 69-76. [DOI: https://dx.doi.org/10.1016/j.ecoleng.2013.06.008]

5. Bennett, E.R.; Moore, M.T.; Cooper, C.M.; Smith, S.; Shields, F.D.; Drouillard, K.G.; Schulz, R. Vegetated agricultural drainage ditches for the mitigation of pyrethroid-associated runoff. Environ. Toxicol. Chem.; 2005; 24, pp. 2121-2127. [DOI: https://dx.doi.org/10.1897/04-357R.1]

6. Needeman, B.A.; Kleinman, P.J.A.; Strock, J.S.; Allen, A.L. Improved management of agricultural drainage ditches for water quality protection: An overview. J. Soil Water Conserv.; 2007; 62, pp. 171-178.

7. Strock, J.S.; Dell, C.J.; Schmidt, J.P. Managing natural processes in drainage ditches for nonpoint source nitrogen control. J. Soil Water Conserv.; 2007; 62, pp. 188-196.

8. Huang, J.; Liu, R.M.; Wang, Q.R.; Gao, X.; Han, Z.Y.; Gao, J.M.; Gao, H.; Zhang, S.B.; Wang, J.F.; Zhang, L. et al. Climate factors affect N₂O emissions by influencing the migration and transformation of nonpoint source nitrogen in an agricultural watershed. Water Res.; 2022; 223, 119028. [DOI: https://dx.doi.org/10.1016/j.watres.2022.119028]

9. Nazaryuk, V.M.; Sidorova, K.K.; Shumny, V.K.; Kalimullina, F.R.; Klenova, M.I. Nitrogen regime of soils with different macrosymbiont genotypes. Eurasian Soil Sci.; 2007; 40, pp. 168-175. [DOI: https://dx.doi.org/10.1134/S106422930702007X]

10. In’t Zandt, D.; Fritz, C.; Wichern, F. In the land of plenty: Catch crops trigger nitrogen uptake by soil microorganisms. Plant Soil; 2018; 423, pp. 549-562. [DOI: https://dx.doi.org/10.1007/s11104-017-3540-2]

11. Nasholm, T.; Kielland, K.; Ganeteg, U. Uptake of organic nitrogen by plants. New Phytol.; 2009; 182, pp. 31-48. [DOI: https://dx.doi.org/10.1111/j.1469-8137.2008.02751.x] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/19210725]

12. Saleem, M.; Fetzer, I.; Harms, H.; Chatzinotas, A. Trophic complexity in aqueous systems: Bacterial species richness and protistan predation regulate dissolved organic carbon and dissolved total nitrogen removal. Proc. R. Soc. B Biol. Sci.; 2016; 283, 20152724. [DOI: https://dx.doi.org/10.1098/rspb.2015.2724] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/26888033]

13. Xiao, H.Y.; Liu, C.Q. Situation of sewage input reflected by nitrogen isotopic composition in a sediment core of Hongfeng Lake. Chin. Sci. Bull.; 2006; 51, pp. 971-976. [DOI: https://dx.doi.org/10.1007/s11434-006-0971-1]

14. Ruiz, G.; Jeison, D.; Rubilar, O.; Ciudad, G.; Chamy, R. Nitrification-denitrification via nitrite accumulation for nitrogen removal from wastewaters. Bioresour. Technol.; 2006; 97, pp. 330-335. [DOI: https://dx.doi.org/10.1016/j.biortech.2005.02.018]

15. Liu, J.K.; Liu, J.L.; Zhang, R.; Zou, Y.Q.; Wang, H.H.; Zhang, Z.M. Impacts of aquatic macrophytes configuration modes on water quality. Water Sci. Technol.; 2014; 69, pp. 253-261. [DOI: https://dx.doi.org/10.2166/wst.2013.573]

16. Chen, L.; Liu, F.; Wang, Y.; Li, X.; Zhang, S.N.; Li, Y.; Wu, J.S. Nitrogen removal in an ecological ditch receiving agricultural drainage in subtropical central China. Ecol. Eng.; 2015; 82, pp. 487-492. [DOI: https://dx.doi.org/10.1016/j.ecoleng.2015.05.012]

17. Zhao, M.M.; Chen, Y.P.; Xue, L.G.; Fan, T.T. Three kinds of ammonia oxidizing microorganisms play an important role in ammonia nitrogen self-purification in the Yellow River. Chemosphere; 2020; 243, 125405. [DOI: https://dx.doi.org/10.1016/j.chemosphere.2019.125405]

18. Ouyang, Q.L.; Gao, G.Q.; Liu, S.S.; Shi, X.L. Study on Characters of Nitrogen Adsorption and Desorption of Sediment in Ganjiang River. Proceedings of the International Conference on Civil, Architectural and Hydraulic Engineering (ICCAHE 2012); Zhangjiajie, China, 10–12 August 2012; 391.

19. Chen, D.Q.; He, H.; Chen, Y.Q. Purification of Nitrogen and Phosphorus in lightly Polluted Landscape River by effective microorganisms combined with submerged plants. Proceedings of the International Conference on Energy Engineering and Environmental Engineering (ICEEEE 2013); Hangzhou, China, 18–19 January 2013; 430.

20. Toyama, T.; Nishimura, Y.; Ogata, Y.; Sei, K.; Mori, K.; Ike, M. Effects of planting Phragmites australis on nitrogen removal, microbial nitrogen cycling, and abundance of ammonia-oxidizing and denitrifying microorganisms in sediments. Environ. Technol.; 2016; 37, pp. 478-485. [DOI: https://dx.doi.org/10.1080/09593330.2015.1074156]

21. Chen, L.A.; Zhang, W.S.; Tan, J.Y.; Shao, X.H.; Qiu, Y.L.; Zhang, F.X.; Zhang, X. Nitrogen and Phosphorus Pollutants Removal from Rice Field Drainage with Ecological Agriculture Ditch: A Field Case. Phyton Int. J. Exp. Bot.; 2022; 91, pp. 2827-2841. [DOI: https://dx.doi.org/10.32604/phyton.2022.024105]

22. Kumwimba, M.N.; Zhu, B.; Muyembe, D.K. Assessing the influence of different plant species in drainage ditches on mitigation of non-point source pollutants (N, P, and sediments) in the Purple Sichuan Basin. Environ. Monit. Assess.; 2017; 189, 267. [DOI: https://dx.doi.org/10.1007/s10661-017-5965-8]

23. Liu, H.S.; Yang, H.; Shang, H.H.; Su, F.L.; Ji, C.Z.; Cheng, J. Effect of the Matrix Dam in the Paddy Field Drainage Ditch on Water Purification Based on the Physical Model Test. Sustainability; 2022; 14, 8620. [DOI: https://dx.doi.org/10.3390/su14148620]

24. Kroger, R.; Hollad, M.M.; Moore, M.T.; Cooper, C.M. Agricultural drainage ditches mitigate phosphorus loads as a function of hydrological variability. J. Environ. Qual.; 2008; 37, pp. 107-113. [DOI: https://dx.doi.org/10.2134/jeq2006.0505] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/18178883]

25. Kroger, R.; Holland, M.M.; Moore, M.T.; Cooper, C.M. Hydrological variability and agricultural drainage ditch inorganic nitrogen reduction capacity. J. Environ. Qual.; 2007; 36, pp. 1646-1652. [DOI: https://dx.doi.org/10.2134/jeq2006.0506] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/17940264]

26. Fan, G.D.; Li, Z.S.; Wang, S.M.; Huang, K.S.; Luo, J. Migration and transformation of nitrogen in bioretention system during rainfall runoff. Chemosphere; 2019; 232, pp. 54-62. [DOI: https://dx.doi.org/10.1016/j.chemosphere.2019.05.177]

27. Zhan, J.; Li, W.J.; Li, Z.P.; Zhao, G.Z. Indoor experiment and numerical simulation study of ammonia-nitrogen migration rules in soil column. J. Groundw. Sci. Eng.; 2018; 6, pp. 205-219.

28. Lv, X.D.; Li, H.B.; Wang, H.; Wang, X.; Li, Y.H.; Ji, X.C.; Yu, Y.Y. Effect of Matrixes and Plants on the Removal of Nitrogen and Phosphorus in Paddy Fields Drainage Ditch. Proceedings of the 2nd International Conference on Renewable Energy and Environmental Technology (REET); Dalian, China, 19–20 August 2014; 516.

29. Cui, N.X.; Zhang, X.; Cai, M.; Zhou, L.; Chen, G.F.; Zou, G.Y. Roles of vegetation in nutrient removal and structuring microbial communities in different types of agricultural drainage ditches for treating farmland runoff. Ecol. Eng.; 2020; 155, 105941. [DOI: https://dx.doi.org/10.1016/j.ecoleng.2020.105941]

30. Ma, Y.H.; Zheng, X.Y.; Fang, Y.Q.; Xu, K.Q.; He, S.B.; Zhao, M. Autotrophic denitrification in constructed wetlands: Achievements and challenges. Bioresour. Technol.; 2020; 318, 123778. [DOI: https://dx.doi.org/10.1016/j.biortech.2020.123778]

31. Fu, J.; Han, J.; Qi, J.; Han, R.; Xu, Y.; Wang, L. Study on the Shortcut Nitrification Process under Low DO and High pH Condition in Normal Temperature. J. Shenyang Inst. Archit. Eng. Nat. Sci. Ed.; 2010; 26, pp. 316-320.

32. Fernando, K.M.C.; Ehoche, O.G.; Atkinson, J.A.; Sparkes, D.L. Root System Architecture and Nitrogen Uptake Efficiency of Wheat Species. J. Agric. Sci.; 2021; 16, pp. 37-53. [DOI: https://dx.doi.org/10.4038/jas.v16i1.9182]

33. Godde, M.; Conrad, R. Immediate and adaptational temperature effects on nitric: Oxide production and nitrous oxide release from nitrification and denitrification in two soils. Biol. Fertil. Soils; 1999; 30, pp. 33-40. [DOI: https://dx.doi.org/10.1007/s003740050584]

34. Qun, Z.; Zhihao, Z.; Ting, Y.; Huchun, X.; Xin, Y.; Jing-jing, X. Effects of COD/N ratios on mainstream simultaneous nitritation and denitritation process. Environ. Ecol.; 2023; 5, pp. 68-73.

35. Wang, J.; Huang, Q.; Li, Y.; Tu, X.S.; Chen, Z.X.; Elrys, A.S.; Cheng, Y.; Ma, L.F. A shift from nitrification to denitrification-dominated N₂O emission in an acidic soil following organic amendment. Biol. Fertil. Soils; 2023; 59, pp. 117-122. [DOI: https://dx.doi.org/10.1007/s00374-022-01680-7]

36. Li, W.H.; Siddique, M.S.; Liu, M.J.; Graham, N.; Yu, W.Z. The migration and microbiological degradation of dissolved organic matter in riparian soils. Water Res.; 2022; 224, 119080. [DOI: https://dx.doi.org/10.1016/j.watres.2022.119080]

37. Tan, X.Z.; Shao, D.G.; Gu, W.Q. Effects of temperature and soil moisture on gross nitrification and denitrification rates of a Chinese lowland paddy field soil. Paddy Water Environ.; 2018; 16, pp. 687-698. [DOI: https://dx.doi.org/10.1007/s10333-018-0660-0]

38. Corre, M.D.; Schnabel, R.R.; Stout, W.L. Spatial and seasonal variation of gross nitrogen transformations and microbial biomass in a Northeastern US grassland. Soil Biol. Biochem.; 2002; 34, pp. 445-457. [DOI: https://dx.doi.org/10.1016/S0038-0717(01)00198-5]

39. Zaman, M.; Chang, S.X. Substrate type, temperature, and moisture content affect gross and net N mineralization and nitrification rates in agroforestry systems. Biol. Fertil. Soils; 2004; 39, pp. 269-279. [DOI: https://dx.doi.org/10.1007/s00374-003-0716-0]

40. Liu, B.; Zeng, Q.; Yan, F.M.; Xu, H.G.; Xu, C.R. Effects of transgenic plants on soil microorganisms. Plant Soil; 2005; 271, pp. 1-13. [DOI: https://dx.doi.org/10.1007/s11104-004-1610-8]

41. Bell, C.W.; Asao, S.; Calderon, F.; Wolk, B.; Wallenstein, M.D. Plant nitrogen uptake drives rhizosphere bacterial community assembly during plant growth. Soil Biol. Biochem.; 2015; 85, pp. 170-182. [DOI: https://dx.doi.org/10.1016/j.soilbio.2015.03.006]

42. Qin, R.Q.; Chen, H. The procession of constructed wetland removal mechanism of pollutants. Proceedings of the 4th International Conference on Mechanical Materials and Manufacturing Engineering (MMME); Wuhan, China, 15–16 October 2016; pp. 568-570.

43. Simek, M.; Santruckova, H. Influence of storage of soil samples on microbial biomass and its activity. Rostl. Vyrob.; 1999; 45, pp. 415-419.

44. Lu, S.; Yang, Y.M.; Yin, H.L.; Su, X.S.; Yu, K.N.; Sun, C. Microbial Community Structure of Arsenic-Bearing Groundwater Environment in the Riverbank Filtration Zone. Water; 2022; 14, 1548. [DOI: https://dx.doi.org/10.3390/w14101548]

45. Kampman, C.; Piai, L.; Hendrickx, T.L.G.; Temmink, H.; Zeeman, G.; Buisman, C.J.N. Effect of temperature on denitrifying methanotrophic activity of ‘Candidatus Methylomirabilis oxyfera’. Water Sci. Technol.; 2014; 70, pp. 1683-1689. [DOI: https://dx.doi.org/10.2166/wst.2014.431] [PubMed: https://www.ncbi.nlm.nih.gov/pubmed/25429458]

46. Wang, C.Y.; Wu, D.F.; Mao, X.M.; Hou, J.M.; Wang, L.; Han, Y.P. Estimating soil ammonium adsorption using pedotransfer functions in an irrigation district of the North China Plain. Pedosphere; 2021; 31, pp. 157-171. [DOI: https://dx.doi.org/10.1016/S1002-0160(20)60054-6]

47. Ma, L.W.; Lindau, C.W.; Hongprayoon, C.; Burhan, W.; Jang, B.C.; Patrick, W.H.; Selim, H.M. Modeling urea, ammonium, and nitrate transport and transformations in flooded soil columns. Soil Sci.; 1999; 164, pp. 123-132. [DOI: https://dx.doi.org/10.1097/00010694-199902000-00007]

48. Fernandes, S.O.; Javanaud, C.; Aigle, A.; Michotey, V.D.; Guasco, S.; Deborde, J.; Deflandre, B.; Anschutz, P.; Bonin, P.C. Anaerobic nitrification-denitrification mediated by Mn-oxides in meso-tidal sediments: Implications for N-2 and N₂O production. J. Mar. Syst.; 2015; 144, pp. 1-8. [DOI: https://dx.doi.org/10.1016/j.jmarsys.2014.11.011]

49. Martinez-Espinosa, C.; Sauvage, S.; Al Bitar, A.; Green, P.A.; Vorosmarty, C.J.; Sanchez-Perez, J.M. Denitrification in wetlands: A review towards a quantification at global scale. Sci. Total Environ.; 2021; 754, 142398. [DOI: https://dx.doi.org/10.1016/j.scitotenv.2020.142398]