Introduction
The global increase in reactive nitrogen (N) fertilizer use has the potential to cause more N loss, posing a significant threat to biodiversity [1]. The use of reactive N fertilizer in agricultural systems is susceptible to losses due to the swift conversion of ammonium (NH4+)-based N fertilizers to nitrate (NO3−N) through microbial nitrification, which has profound implications for human health and the environment. Other forms of reactive N loss include ammonia (NH3) volatilization and nitrous oxide (N2O) gas emission through nitrification–denitrification pathways, leading to the emission of the ozone-depleting greenhouse gas nitrous oxide (N2O) [2]. Consequently, there has been a global initiative to develop fertilizer management strategies that optimize N application and minimize N loss [3]. Current strategies include optimizing the rate and timing of N fertilizer application, implementing crop rotations, and using enhanced fertilizer formulated with inhibiting compounds or inhibitors. The use of nitrification inhibitors has often proven to be an effective strategy to reduce N losses and thereby increase N fertilizer efficiency, particularly for ammonium-based fertilizers. Nitrification inhibitors delay the conversion of NH4+-N to NO3−N and decrease N loss by inactivating the ammonia monooxygenase (AMO) enzyme [3], limiting the first-rate step of autotrophic nitrification carried out by ammonia-oxidizing archaea (AOA), ammonia-oxidizing bacteria (AOB), and Comammox bacteria [4]. However, their use has often been linked to environmental contamination, non-target effects on beneficial soil microbes [5], limited efficacy dependent on soil conditions (moisture, soil pH, organic matter, temperature) [6], and the risk of NH3 volatilization [7, 8] following NH4+ accumulation.
Soil pH is particularly important as it is a key factor influencing nitrification processes in soils by affecting the concentration and availability of substrates [9] for microbial N cycling and influencing bacterial diversity and community structure [10]. Soil pH has been shown to be the primary factor driving changes in the community of nitrifying bacteria (AOA, AOB, and Comammox) across a soil pH gradient. For example, AOB was reported to dominate and functionally regulate ammonia oxidation in neutral and alkaline environments, whereas AOA and Comammox bacteria prefer to regulate ammonia oxidation in acidic environments [4, 11]. Given the challenges associated with the application of nitrification inhibitors and the economic cost, which could limit its application in resource-poor agricultural systems, it is necessary to explore alternative and easily sourced stable soil amendments that can slowly release N and enhance N use efficiency in cropping systems.
Biochar has been proposed as a promising candidate for N retention and NH3 volatilization mitigation following soil amendment. However, the performance of pristine biochar in the sorption of cations such as (NH4+) or anions (NO3−) could be complex and limited by pyrolysis conditions, inherent characteristics such as surface charges, C/N ratio, labile nutrients, functional groups, and sorptive properties [12]. Therefore, it is crucial to modify and functionalize the surface of pristine biochar to enhance its adsorption affinity between bare biochar and ions. Ball-milling is of particular interest; this technology is a powerful non-equilibrium physico-mechanical treatment that mechanically reduces the grain size of biomass to nanoscale [13]. Few studies have utilized ball-milling technology in the production of engineered biochar with enhanced physiochemical properties [13–15]. The mechano-physical treatment of biochar via ball-milling yields nano-biochar characterized by reduced particle size, increased internal and external surface area for ion sorption, and augmented acidic surface functional groups [13, 16]. The proposed mechanisms for nitrogen (N) retention and ammonia (NH3) volatilization mitigation induced by ball-milled biochar include increased soil pH buffering, ammonium (NH4+) and nitrate (NO3−) capture on its extensive internal and external surface area and pore spaces [17], oxygen-containing functional groups due to short- and long-term oxidation [18, 19], sorption due to ion exchange, NH4+ sorption via chemisorption ammonia fixation, ion exchange with columbic forces or an association with sulfur-functional groups [20, 21], and alteration of surface functional groups. However, current research on the application of ball-milling technology in biochar modification, particularly for N retention in soil systems, remains scarce. To achieve effectiveness at scale, biochar often necessitates high application rates, e.g., 20–100 t ha−1 [22]. Therefore, it is worthwhile to explore strategies to reduce biochar application rates without compromising its effectiveness, thereby increasing its affordability and acceptance as a soil amendment technique for N retention in agricultural systems.
In the tropics, soil amendment with neem (Azadirachta indica) seed cake has been shown to inhibit microbial nitrification [23, 24] by suppressing the nitrifying bacteria involved in the process due to the release of bioactive compounds and antimicrobial properties (azadirachtin). [25, 26] confirmed the potent antibacterial activity of neem materials against various types of bacteria. [27] also reported the efficacy of neem seed cake and oil in inhibiting the nitrification of urea fertilizer. In addition to nitrification inhibition efficacy, neem seed cake possesses fertilizing potential due to its nutrient-rich constituents [28]. Given the large application rate often required for biochar success at field scale and the competing need for feedstock, a more sustainable synergy could be achieved by combining reduced ball-milled biochar and neem seed cake rather than applying each separately. When combined, the adsorption capacity of ball-milled biochar and the antimicrobial effects of neem seed cake can work synergistically to retain N and control nitrification processes. Besides N retention, this combination can also provide other soil ecosystem benefits such as pH buffering, soil biology activation, increased soil organic carbon storage, improvement of soil physical properties, and act as a slow-release fertilizer for crops. A significant knowledge gap still exists regarding the nitrification inhibition efficacy and potential NH3 volatilization mitigation of ball-milled biochar-neem seed cake co-amendment. Therefore, studying the impact of engineered biochar-organo amendment could provide knowledge on its application in resource-poor systems, and the identification of appropriate soil types based on microbial community sensitivity. This knowledge will enable a more targeted and effective use of this novel approach towards enhancing soil N retention while minimizing environmental losses in soil systems. In this study, we aim to: (i) establish the impact of ball-milled biochar, neem seed cake, and the co-application of ball-milled biochar-neem seed cake on nitrification dynamics and NH3 volatilization in three contrasting soil types (varying pH levels) and; (ii) to determine the effects of ball-milled biochar, neem seed cake and their co-application on the abundance of nitrifying bacteria communities (ammonia oxidizers and comammox). We hypothesized that the ball-milled biochar-neem seed cake combination will achieve nitrification inhibition and NH3 volatilization mitigation by various mechanisms such as NH4+-N sorption, alteration of nitrifying microbial communities, modification of soil physico-chemical properties, and restricting microbial nutrient substrate availability irrespective of differences in soil type.
Materials and methods
Collection and preparation of soil and substrates
Surface soil samples (0–15 cm) were collected from three sites: two conventional agricultural fields and one secondary forest, covering an area of 0.25 ha each. The sites were located in Nigeria (NG) and Egypt (EG) with contrasting soil textures and pH values. The specific locations were: (i) Akungba-Akoko, Nigeria (7.48°N, 5.75°E): Secondary forest with sandy loam soil; (ii) Ikoro-Ekiti, Nigeria (7.83°N, 5.03°E): Cropland with sandy clay loam soil; (iii) Sohar, Egypt (24.15°N, 56.47°E): Cropland with fine sand soil. Akungba-Akoko is situated in the deciduous rainforest in south-west Nigeria, with a vegetation type that reflects rainforest and guinea savannah. The long-term average annual rainfall at Akungba-Akoko is 1318 mm, out of which about 85% falls between June and September, with the remainder from January to May. Average air temperatures range between 23.3 and 24.7 °C. The soil from Akungba-Akoko is of the order Haplic Lixisols. Lixisols are considered marginal lands that are mildly acidic and increase in clay content with depth, and depositional spots can have an enrichment of base cations through aeolian deposits [29]. The soil from Ikoro-Ekiti exhibits a tropical wet and dry or savanna climate. The long-term average annual rainfall at Ikoro-Ekiti is 148.18 mm, and the average air temperature ranges between 22.0 and 38.9 °C. The soil at Ikoro-Ekiti is of the order Plinthosol-Lixisol [29]. Soil from Egypt is a calcareous alkaline soil from agricultural cropland with cereal-vegetable crop rotation history in the Arab El-Awamer at Assiut Governorate, Upper Egypt with an average annual temperature of 26 °C and 110 mm of average annual precipitation. The soil order is characterized as Entisols [29]. The collected soil was air-dried, thoroughly mixed, and ground to pass through a 2-mm sieve before being stored in a refrigerator prior to the incubation experiment. Soil texture was determined based on wet-sieving and sedimentation, according to [30].
Pristine biochar was produced in a reactor by combustion of wood chips at 500 °C for 75 min under no‑oxygen condition. Thereafter, produced biochar was mechanically modified by ball-milling in a planetary ball mill to produce very fine particles (nanoparticle) following protocols of [13]. Briefly, woodchip biochar was placed inside each agate container (500 mL) with agate balls of diameter size 6 mm and weight of 180 g in an XQM 2A ball mill machine and operated at 300 rpm with the direction of rotation changed every 30 min for 7 h. On completion, collected biochar were labelled ball-milled biochar (BM-Biochar). Neem seed cake was purchased as a commercially available substrates from an agro-shop. The bioactive ingredient (azadirachtin) concentration in neem seed cake was determined by methanolic extraction and filtrates were analysed using a 6400 series Triple Quad Liquid Chromatography Mass Spectrometer (LC/MS) and a MassHunter WorkStation software (Agilent Technologies, Santa Clara, CA, USA). Air-dried samples of soil and neem seed cake were ball-milled and analyzed for total C and N content by dry combustion using a VarioMax® CHN analyzer (Elementar Analysesysteme GmbH, Langenselbold, Germany). Total P was determined using an ICP-OES. Dissolved organic C (DOC), dissolved N (DN), and dissolved P (DP) were extracted in deionized water (dH2O) (1:10 sample: dH2O ratio) and analyzed using a TOC-LCPH/CPN analyser (Shimadzu) and spectrophotometry according to [31]. Sample pH was assessed in a 1:10 sample:dH2O suspension using a glass electrode. The chemical properties of the soil and organic substrates are presented in Tables S1 and S2.
Microcosms experiment
An incubation experiment was conducted using microcosms. 1 kg of air-dried soil from each of the three soil types was placed in a microcosm PVC plastic column (17 cm in diameter and 12 cm in height) at 25 °C and 60% water holding capacity (WHC). Samples were initially pre-wetted and incubated for 2 weeks under 30 °C at 60% WHC to equilibrate the soil before the application of treatments. Ammonium chloride (NH4Cl) fertilizer was applied to all incubated soils/treatments at the rate of 0.5 g N kg−1 dry soil. Four treatments with five replicates each were assigned to three soil types (neutral, acidic, and alkaline) in a completely randomized design. The treatments were as follows: (a) Control [NH4Cl only] (CK), (b) neem seed cake, (c) BM-biochar, and (d) BM-biochar + neem seed cake (1:1w/w), all applied at a modest rate of 2% w/w of dry soil. Treatment combinations gave a total of 60 microcosms. Soil samples were thoroughly agitated with corresponding amendments, including the control and all plastic columns were incubated at room temperature at 32 ± 1 °C for 30 days. Deionized water was regularly added to maintain WHC of the soil at 60%. During the incubation, triplicate soil samples were repeatedly extracted from all treatments for NH4+ and NO3− concentration on days 0, 7, 14, 21, and 28 with 2 M KCl (soil-to solution ratio 1:5) by shaking for 1 h. The extracts were filtered through Whatman number 42 filter papers and analysed for mineral N (NH4+ and NO3−) using a continuous flow analyzer (AA III, Norderstedt, Germany). All incubation jars were destructively harvested at the end of the experiment (day 30).where: Final NO3− is the nitrate concentration at the end of incubation period.
Initial NO3− is the nitrate concentration at the beginning of the incubation period.
Days of incubation is the duration of the experiment in days.
The percentage of nitrification inhibition was determined following [2] protocol by calculatingwhere; A = Nitrification rate of soil without amendment (control).
B = Nitrification rate of soil with amendment.
Soil ammonia (NH3) volatilization
NH3 volatilization was measured from three replicates per treatments by the venting method [33]. Inside the microcosm chamber, two circular pieces of sponge with a thickness of about 2 cm were evenly soaked with glycerophosphate (15 mL of 5% phosphoric acid in 4% of glycerol solution) and placed in each plastic column. The lower sponge is 4 cm from the bottom of the column and the upper sponge is horizontal with the top of the column. The upper sponge was to absorb NH3 gas from the air and prevent contamination, while the lower sponge traps NH3 gas volatilized from soil in the column. When sampling, the lower sponge was taken out, transferred into a plastic bag, and replaced with a freshly soaked glycerophosphate sponge. The upper protective sponge was replaced at an interval of 7 days. The removed lower sponges were placed into 500 mL plastic bottles respectively, and 300 mL 1 mol·L−1 KCl solution was added to completely immerse the sponges. After one hour of oscillation, the N concentration in the solution was analyzed by a continuous flow analyzer (AA III, Norderstedt, Germany). NH3 gas samples were collected from at an interval of 7 days for 28 days of soil incubation. Cumulative NH3 volatilization was recorded as the sum of daily NH3 gas fluxes from day 1 until day 28.
DNA extraction and quantitative PCR
After 28 days of incubation, soil DNA was extracted from 0.5 g soil following the protocols of FastDNA® SPIN Kit for Soil (MP Biomedicals, USA). DNA concentration was detected by NanoDrop 2000 UV–Vis spectrophotometer (Thermo Scientific, USA), and DNA quality was detected by 1% agarose gel electrophoresis. The soil nitrifying microorganism AOA amoA gene was amplified using the primer sequences A26F (5′-GACTACATMTTCTAYACWGAYTGGGC-3′) and A416R (5′-GGKGTCATRTATGG WGGYAAYGTTTGG-3′) [34]. The specific genus and species targeted for AOA using the amoA gene, which encodes the ammonia monooxygenase enzyme include Nitrosopumilus—a well-known representative of AOA, commonly found in marine environments and soils, Nitrososphaera—often found in terrestrial environments, Nitrosotalea—commonly dominant in acidic soils, and Nitrosocaldus—often found in hot spring environments. The primer sequence amoA-1F (5′-GGGGTTCTACTGGTGGT-3′) and amoA-1R (5′-CCCCTCKGSAAAGCCTTCTTC-3′) were used to amplify the AOB amoA gene [35]. Specific genus and species targeted for AOB include Nitrosomonas—prevalent in soil environment, and Nitrosospira—often found in soils and freshwater environments. The primer sequence ComamoA F (5′-AGGNGAYTGGGAYTTCTGG-3′) and ComamoA R (5′-CGGACAWABRTGAABCCCAT-3′) were used to amplify ComamoA [36]. Genus and species targeted for Comammox include Nitrospira—capable of complete nitrification, converting ammonia directly to nitrate. The PCR reaction procedure was performed as follows: 1 μL of 10 μM upstream primer and 1 μL downstream primer, 1 μL of SYBR® PremixExTaq™ II (TliRNaseHPlus) ROXplusL, 25 μL of 2 × TaqMasterMix and 22 μL of water, a total of 50 μL. The PCR amplification conditions of the amoA gene were as follows: pre-denaturation at 94 °C for 5 min, 30 cycles of 94 °C for 30 s, 55 °C for 30 s, 72 °C for 30 s, and extension at 72 °C for 10 min after the last cycle. Each sample was replicated three times, and a standard curve was established with known copy numbers of soil-nitrifying microorganisms using a dilution series of linearized pGEM-T Easy plasmids (Promega) containing clones of each PCR fragment.
Statistical analysis
Two-way analysis of variance using General Linear Model (GLM) was deployed to determine interactions between soil amendments and soil types on the dynamics of NH4+ and NO3−release, NH3 volatilization, soil properties and gene copy numbers of nitrifying bacteria. Prior to analysis and where necessary, dependent variables were log-transformed to satisfy the assumptions of normality and homoscedasticity. When there were no significant interactions, one-way analysis of variance was then used to analyze differences in the tested parameters amongst treatments. Tukey’s HSD post hoc test at the 5% probability level was used to test the significance of differences between treatment means. Pearson correlation analysis was performed using SPSS to explore the relationship between soil chemical properties and the abundance of nitrifying soil bacteria. Principal component analysis (PCA) was also conducted using PAST 4.16c to determine the effect and relationship of soil amendments on the nitrifying bacteria abundance across soil types (Fig. 1).
Fig. 1 [Images not available. See PDF.]
a–c Boxplots of comparative effect of neem seed cake (NSC), ball-milled biochar (BM BC), and ball-milled biochar + neem seed cake (BMBC + NSC) on the percentage of nitrification inhibition of neutral, alkaline, and acidic soil after 28 days of incubation. Different lowercase letters on bars indicate significantly different means at p < 0.05 based on the Tukey’s HSD test
Results
Dynamics of soil ammonium N (NH4+-N)
We observed no significant interaction (p < 0.05) between amendments and soil types on soil ammonium-N (NH4+-N) concentrations (Table S3). However, the main effects of amendments and soil types were individually significant (p < 0.05) (Fig. 2). NH4+-N contents varied throughout the incubation period, primarily due to differences in soil pH across soil types. The contents of NH4+-N differed over the incubation period due to the differences in soil type (pH). In the neutral pH soil, NH4+-N ranged from 9.4 to 63.2 mg kg−1 from day 0 to day 28 across all treatments (Fig. 2a). The control treatment (NH4Cl) without soil amendment addition showed a steady decline in NH4+-N content from day 0 to day 28 in the neutral pH soil, while the BM-biochar, neem seed cake and BM-biochar + neem seed cake showed opposite trend by accumulating NH4+-N content than CK (Fig. 2a). After 28 days of incubation, BM-biochar + neem seed cake, neem seed cake and sole BM-biochar treatment respectively inhibited N nitrification and accumulated 43%, 25% and 35% of NH4+-N content more than CK in the neutral pH soil. NH4+-N content accumulation in the neutral pH soil was in the order of BM-biochar + neem seed cake ˃ BM-biochar ˃ neem seed cake ˃ CK (Fig. 2a). For the alkaline soil, NH4+-N content ranged from 24.8 to 78.4 mg kg−1 from day 0 to day 28 across all treatments (Fig. 2b). The CK showed a peak of NH4+-N at day 7, and thereafter displayed a steady decline of NH4+-N from day 14 to day 28. NH4+-N content accumulation wasn’t significantly (p < 0.05) different between BM-biochar + neem seed cake, neem seed cake and BM biochar from day 0 until day 21. At day 28, sole BM-biochar and CK accumulated the lowest NH4+-N content (Fig. 2b). NH4+-N content accumulation in the alkaline soil was in the order of BM-biochar + neem seed cake ˃ neem seed cake ˃ BM-biochar ˃ CK (Fig. 2b). In the acidic soil, NH4+-N content accumulation followed a trend of BM-biochar ˃ neem seed cake ˃ CK ˃ BM-biochar + neem seed cake after 28 days of incubation (Fig. 2c). NH4+-N content was generally within the range of 31.2–78.4 mg kg−1. Overall, BM-biochar + neem seed cake had accumulated more NH4+-N content particularly in the neutral and alkaline soil (Fig. 2a-c).
Fig. 2 [Images not available. See PDF.]
Dynamics of ammonium nitrogen (NH4+-N) and nitrate nitrogen (NO3−-N) release in the neutral (a, d), alkaline (b, e), and acidic soil (c, f) affected by neem seed cake, ball-milled biochar, and ball-milled biochar + neem seed cake over 28 days of soil incubation
Dynamics of soil nitrate (NO3−-N)
Statistical analysis showed no significant (p < 0.05) interaction of soil amendment × soil types on soil nitrate N (NO3−-N) (Table S3). NO3−-N accumulation indicated nitrification was significantly higher in CK treatment but followed a quadratic release pattern across all soil types (neutral, alkaline, and acidic) (Fig. 2d–f). In the neutral soil, NO3—N content ranged between 9.2 and 65.8 mg kg−1, alkaline; 11.5–75.6 mg kg−1, and acidic soil; 15.4–59.6 mg kg−1 (Fig. 2d–f). The nitrification of NH4+-N to NO3−-N was generally inhibited by amendments across treatments except in the CK soil. In reverse to NH4+-N accumulation, NO3−N accumulation was in the order of CK ˃ BM-biochar ˃ neem seed cake ˃ BM-biochar + neem seed cake in the neutral soil, CK ˃ BM-biochar ˃ neem seed cake ˃ BM-biochar + neem seed cake in alkaline soil and in acidic soil; CK ˃ BM-biochar + neem seed cake ˃ neem seed cake ˃ BM-biochar after 28 days of incubation (Fig. 2d–f).
Soil nitrification inhibition rate
The inhibition of nitrate production varied with soil type and soil amendment. Soil amendments generally inhibited nitrate production at an appreciable rate (Fig. 1). In the neutral soil, the percentage of nitrification inhibition rate ranged between 61 and 75%, 61–70% in the alkaline soil, and 10–70% in acidic soil over 28 days of incubation. An interesting finding was observed in the acidic soil, where BM-biochar + neem seed cake had significantly (p < 0.05) decreased the percentage inhibition of NO3-N compared to BM-biochar and neem seed cake (Fig. 1c).
Soil mean nitrification rate
In Fig. 3, we present the overall soil mean nitrification rate after 28 days of soil incubation. There was no significant (p < 0.05) interaction of soil amendment × soil types on mean nitrification rate (Table S3). However, across soil type and treatment, we recorded the highest mean nitrification rate (p < 0.05) in CK.
Fig. 3 [Images not available. See PDF.]
Effect of neem seed cake (NSC), ball-milled biochar (BM BC), control (CK), and ball-milled biochar + neem seed cake (BM-BC + NSC) on the mean nitrification rate of neutral, alkaline, and acidic soil after 28 days of incubation. Bars represent standard error (n = 4). The different lowercase letters above the figures indicate significant differences between different treatments within the same soil at p < 0.05 by Tukey’s HSD test; the different capital letters above the figures indicate significant differences between different soils in the same treatment at p < 0.05 by Tukey’s HSD test
Soil ammonia (NH3) Volatilization
The dynamics of the rate of NH3 volatilization after 28 days of incubation are presented in Figs. 4 and 5. Our statistical model found non-significant (p < 0.05) interaction of soil amendments × soil types on NH3 volatilization (Table S3). Across incubated soil types, cumulative NH3 volatilization was significantly (p < 0.05) lowest in the acidic soil, while the highest volatilization occurred in the alkaline soil, which ranged between 16.55 and 46.29 mg m−2 h−1 (Fig. 5). CK showed a linear decline of NH3 volatilization from day 1–28 in all soil types. In the neutral and alkaline soil, soil amendments were not significantly (p < 0.05) different from each other except for CK, and all amendments volatilized more NH3 gas than CK (Fig. 4a and b). Furthermore, CK showed a linear decline in NH3 volatilization from day 1–day 28 across all soil types. In the alkaline soil, BM-biochar + neem seed cake treatment showed an increasing pattern of NH3 volatilization, while a reverse trend occurred in the acidic soil. In the acidic soil, sole neem seed cake and BM-biochar volatilized more cumulative NH3 gas than CK (Fig. 5).
Fig. 4 [Images not available. See PDF.]
Dynamics of the rate of NH3 volatilization of neem seed cake, ball-milled biochar, and ball-milled biochar + neem seed cake in the neutral (a), alkaline (b), and acidic soils (c) under each sampling day after 28 days of incubation. Bars represent standard error (n = 4)
Fig. 5 [Images not available. See PDF.]
Cumulative NH3 volatilization of control (CK), neem seed cake, ball-milled biochar (BM Biochar), and ball-milled biochar + neem seed cake (BM Biochar + Neem) in the acid, neutral, and alkaline soils after 28 days of incubation. Bars represent standard error (n = 4). The different lowercase letters above the figures indicate significant differences between different treatments within the same soil at p < 0.05 by Tukey’s HSD test; the different capital letters above the figures indicate significant differences between different soils in the same treatment at p < 0.05 by Tukey’s HSD test
Abundance of soil nitrifying microorganisms
The interaction effect between soil amendments and soil type on the abundance of soil amoA gene copy number in nitrifying bacteria was not statistically significant (p < 0.01, Fig. 6). For ammonia-oxidizing archaea (AOA), the application of BM-biochar + neem seed cake resulted in a two-fold increase in abundance in both neutral and acidic soil types, while causing a 50% decrease in alkaline soil compared to the control (CK) (Fig. 6a). A similar trend was observed in ammonia-oxidizing bacteria (AOB), where BM-biochar + neem seed cake treatment led to a two-fold increase in abundance in both neutral and acidic soil types, while inducing a 50% decrease in alkaline soil compared to the control (Fig. 6b). The abundance of complete ammonia oxidizers (comammox) was 1.6 times higher than the control in neutral and acidic soils, and 1.4 times higher in alkaline soil when treated with BM-biochar + neem seed cake (Fig. 6c). Principal Component analysis (PCA) was deployed to evaluate the effect of amendments on nitrifying bacteria composition across soil types (Fig. 7). In the neutral soil, Principal component 1 (PC1) explained 90.71%, while principal component 2 (PC2) explained 8.22% of the total variance (Fig. 7a). Comammox bacteria had a strong positive loading on PC1, while AOB and AOA had a negative loading. On PC2, AOA, AOB and Comammox bacteria had positive loadings. In the alkaline soil, PC1 explained 82.03%, while PC2 explained 15.27% of the total variance (Fig. 7b). Comammox bacteria and AOA had a strong positive loading on PC1, while AOB had a negative loading. On PC2, AOA, AOB, and Comammox bacteria had positive loadings. In the acidic soil, PC1 explained 87.13%, while PC2 explained 8.92% of the total variation. Comammox bacteria had a strong positive loading on PC1, while AOB and AOA had a negative loading. On PC2, AOA, AOB and Comammox had positive loadings (Fig. 7c). As shown by PCA biplot, Comammox bacteria was clustered in the neem seed cake ellipsis distant from other treatments, AOB and AOA were clustered in BM-biochar + neem seed cake, BM-biochar and CK ellipsis in the neutral soil (Fig. 7a). In the alkaline soil, Comammox bacteria was clustered in the neem seed cake ellipsis, AOB and AOA was clustered in BM-biochar + neem seed cake, neem seed cake and CK (Fig. 7b). In the acidic soil, Comammox bacteria was clustered in the neem seed cake and control (CK) ellipsis, AOB in the BM-biochar and BM-biochar + neem seed cake ellipsis, while AOA was clustered in treatments suggesting a spread across all treatments (Fig. 7c).
Fig. 6 [Images not available. See PDF.]
Abundance of amoA genes of AOA (a), AOB (b), and Comammox bacteria (c) after 28 days of incubation with soil amendments in neutral, alkaline, and acidic soils. The different lowercase letters above the figures indicate significant differences between different treatments within the same soil at p < 0.05 by Tukey’s HSD test; the different capital letters above the figures indicate significant differences between different soils in the same treatment at p < 0.05 by Tukey’s HSD test
Fig. 7 [Images not available. See PDF.]
Principal component analysis score and biplot of nitrifying bacteria communities affected by control (CK), neem seed cake (NSC), ball-milled biochar (BM BC), and ball-milled biochar + neem seed cake (BM BC + NSC) addition after 28 days of soil incubation
Soil pH, total C, and total N after incubation period
After 28 days of incubation, soil amendments, soil type, and their interactions significantly influenced soil pH, total carbon (C), and total nitrogen (N) content (Table S4). In neutral soil, the application of BM-biochar alone and BM-biochar combined with neem seed cake resulted in significant increases in soil pH by 1.94 and 0.19 units, respectively. Conversely, neem seed cake application decreased the neutral soil pH by 0.95 units compared to the control (CK). Both BM-biochar and BM-biochar + neem seed cake treatments increased total C content by 48%. Additionally, the BM-biochar + neem seed cake treatment increased total N content by 6% compared to the control (p < 0.001, Table S4). Neem seed cake and BM-biochar + neem seed cake decreased soil pH by 1.21 and 0.16 units respectively in the alkaline soil. In contrast, BM-biochar increased soil pH by 0.07 unit in comparison to CK. Total C was significantly higher in BM-biochar and BM-biochar + neem seed cake by 44% and 47% respectively in alkaline soil. In the acidic soil, BM-biochar + neem seed cake surprisingly caused a decrease in soil pH by 0.06 unit while BM-biochar application induced an increase by 1.5 unit in comparison to CK. Total C and N were the highest in BM-biochar and BM-biochar + neem seed cake by 36%, while neem seed cake increased total N by 10% in comparison to CK.
Relationship between inorganic N dynamics, ammonia volatilization (NH3), nitrification rate, microbial abundance, and soil chemical properties
The Pearson correlation analysis between the abundance and nitrifying activity of the three nitrifying bacteria and soil chemical properties under different amendments showed that the abundance of AOA was significantly negatively correlated with SOM, NH4+-N and NO3−-N, the abundance of AOB was significantly positively correlated with SOM, NO3—N and nitrification rate (Table S5). The abundance of Comammox bacteria was significantly positively correlated with total N, NO3—N, SOM and nitrification rate but negatively correlated with soil pH. Nitrification rate showed a significant positively correlation with soil pH and NO3—N but negative correlation with NH4+-N. Cumulative NH3 gas volatilization showed a significantly positively correlation with soil pH, and slightly negative correlation with soil NO3—N. SOM was significantly positively correlated with total N, NH4+-N and NO3—N. Soil pH was significantly negatively correlated with NH4+-N and NO3—N.
Discussion
NH4+-N and NO3−N dynamics
In this study, we observed a decrease in NO3−N production and an accumulation of NH4+-N content over a 28-day period in incubated microcosm soils. This followed the amendment of BM-biochar and neem seed cake (Fig. 2), an outcome that aligns with the known effects of pristine biochar on N retention and transformation. Prior studies [37–39] have often linked the impact of pristine biochar on soil NO3−N reduction and NH4+-N accumulation to factors such as an increased soil C/N ratio, direct sorption of NH4+ on the biochar surface, and the release of nitrifying bacteria inhibiting compounds like terpenes and α–pinene [40]. These factors collectively inhibit N nitrification. However, our study attributes the accumulation of NH4+-N and the release of NO3−N to a complex interaction between soil amendment properties, soil pH, the abundance of the nitrifying microbial community, and nutrient cycling processes. In neutral and alkaline soil (Fig. 2a and b), the amendment of BM-biochar and neem seed cake resulted in a greater accumulation of NH4+-N ions compared to other treatments. We attribute this to the ball milling technique used to optimize pristine wood chip biochar, which likely led to increased NH4+-N sorption and accumulation due to an enhanced biochar surface area, pores, and functional groups [41]. Our findings contrast with previous reports that indicate a decrease in the direct sorption of NH4+ ions in soils by pristine biochar pyrolyzed at high temperatures (≥ 500 °C), due to a reduction in microbial substrate [42]. This contrast underscores the improved physicochemical and sorptive properties of ball milled biochar compared to pristine biochar. Soil pH plays a significant role in N transformation, influencing gross mineralization [43], net immobilization of NH4+ and NO3−N [43, 44], nitrification rates [45] and dissimilatory nitrate reduction to ammonium [46]. In a turn of pH-dependent effects, BM-biochar induced an increase in soil pH towards alkalinity, leading to an increased NO3−N content in both neutral and alkaline soil (Fig. 2d and e). This is consistent with the general association of NO3−N release and nitrification activity with high pH soils [47], a correlation further affirmed by a Pearson correlation analysis showing significant correlations between soil pH and NO3−N and NH4+ (Table S5). The addition of neem seed cake decreased the pH of neutral and alkaline soil (Table S4), a change we attribute to the decomposition and release of its inherent acidic organic compounds and organic N into the soil solution. The acidifying effect of neem seed cake has been previously demonstrated by [48]. Despite the decrease in soil pH, neem seed cake accumulated more NO3−N than BM-biochar + neem seed cake treatments, likely due to heterotrophic nitrification. Heterotrophic nitrification is common in agricultural soils within the pH range of 4.0–7.4 and plays a crucial role in maintaining soil inorganic N balance [49, 50]. Contrary to buffering pH changes, BM-biochar + neem seed cake amplified and stimulated NO3−N content in the acidic soil (Fig. 2c, f and Table S4). A possible explanation for this could be enhanced heterotrophic nitrification and an altered nitrifying microbial community under acidic conditions. Previous biochar studies have shown that heterotrophic nitrification is dominant in strongly acidic soil and that the rate of heterotrophic nitrification increases as pH value decreases [51].
NH3 volatilization
We observed a quadratic increase in NH3 volatilization in both neutral and alkaline soil over time, following the co-amendment of BM biochar and neem seed cake. This was also observed in acidic soil treated solely with neem seed cake and BM biochar, due to the accumulation of NH4+ [52]. The increase in NH3 volatilization due to soil amendment was attributed to the maintenance of the pH range within critical levels and the activity of microbial communities required to convert NH4+ to NH3 [53]. In neutral and alkaline soil, BM biochar and neem seed cake may have synergistically affected soil properties and microbial processes by improving soil structure, aeration, water retention, and nutrient sorption, thereby creating a favorable environment for microbial activity. Collectively, these factors may have modulated microbial N transformations and NH3 volatilization. NH3 volatilization is a soil physicochemical process sensitive to a pH range of 7.6 – 8, with emissions increasing within this range [54]. Furthermore, our Pearson correlation analysis demonstrated a significantly positive correlation between soil pH and NH3 volatilization (Table S5). Despite a 0.25 unit decrease in soil pH in neutral soil co-amended with BM biochar and neem seed cake (from 7.32 to 7.07), and a 0.55 unit decrease in alkaline soil (from 8.64 to 8.09) at the end of incubation, these pH ranges were sufficient to increase volatilization due to the accumulation of NH4+-N. This aligns with previous reports [55] that NH3 emission can occur in soils with a pH as low as 5.5 when a large amount of NH4+ accumulates [56]. The highest NH3 volatilization was found in alkaline soil, indicating that NH3 volatilization is a major N loss pathway in alkaline soil [57]. Clearly, the magnitude of NH3 volatilization was affected by soil pH [58]. This suggests that lower nitrification rates may reduce soil acidity, as experienced with BM biochar in acidic soil, but pH increase by BM biochar could still result in amplified NH3 volatilization [59]. Interestingly, despite a decrease in pH caused by the addition of neem seed cake in acidic soil, NH3 volatilization increased. We attribute this to several factors, such as the alteration of the microbial community (an increase in microbial organic N users compared to inorganic N users), leading to increased mineralization and nitrification processes, and subsequently increasing the amount of NH4+ available for transformation to NH3. In addition, neem seed cake may have influenced NH3 volatilization by altering soil chemistry through the increase of soil EC and exchangeable cations. High levels of exchangeable cations can displace NH4+ from exchange sites, making it more susceptible to volatilization [60]. However, while neem seed cake may have decreased soil pH, which typically should reduce NH3 volatilization, the overall effect on NH3 volatilization is complex and could be influenced by soil properties and interactions [61].
Changes in soil pH, Total C and N after incubation
BM-biochar and a combination of BM-biochar and neem seed cake demonstrated contrasting effects on soil pH, varying by soil type. BM-biochar alone increased and buffered pH changes in neutral and acidic soil. In contrast, the combination of BM-biochar and neem seed cake, as well as neem cake alone, resulted in a decrease in soil pH in the acidic soil (Table S4). The primary cause of this soil acidification can be attributed to soil chemistry, specifically the decomposition and release of acidic organic compounds from neem, organic nitrogen nitrification, and nitrogen fertilizer reactions, which released H+ ions for each mole of NH4+ introduced into the soil solution [62]. This finding contradicts previous reports of increased pH in sandy savannah soil and greenhouse trials following neem seed amendment [50, 63]. Conversely, the alkalinity of BM-biochar increased soil pH. The significant increase in total C by BM-biochar and the combination of BM-biochar and neem seed cake, compared to neem seed cake alone, could be attributed to the addition of intrinsic carbon from biochar. Neem seed maintained total C levels comparable to baseline levels after 28 days of incubation. This observation aligns with findings reported by [50], who suggested that biologically active ingredients in neem seeds, such as azadirachtin, di- and tri-terpenoids, pentanoterpenoids, and non-terpenoidal compounds, are toxic to microbial biomass [64]. They also noted that it takes several weeks, beyond the 28 days observed in our study, for microbial adaptation to the soil-neem seed cake environment before humification can occur. The slight increase in total N in the acidic soil due to neem seed cake could be attributed to the release of organic N following gradual decomposition [28].
Soil amendment influence nitrifying microbial communities through altered pH and nitrogen availability
The abundance of AOA, AOB, and Comammox bacteria was influenced by changes in soil pH and N availability due to the application of BM biochar, neem seed cake, and their combination. This was confirmed by the correlation between AOB and Comammox bacteria with soil pH, and AOA, AOB, and Comammox bacteria correlation with NH4+ and NO3− (Table S5). In neutral, alkaline, and acidic soils, AOB was predominantly found, followed by AOA in soils amended with BM-biochar, BM biochar + neem seed cake, and control (CK). In contrast, Comammox bacteria were dominant in soils treated with sole neem seed cake and the control (CK) treatment in acidic soil (Figs. 6 and 7). These differences can be attributed to variations in soil pH. The increased soil pH due to BM biochar, and the buffered soil pH due to BM biochar + neem seed cake, may have led to the abundance and activity of AOB while suppressing AOA and Comammox bacteria. However, our observations contrast with the findings of [65]. Numerous studies [66–68] have demonstrated that soil pH is a crucial factor affecting the abundance of AOA, AOB, and Comammox bacteria in soil systems. AOA prefers acidic soil with low NH4+ availability, while neutral and alkaline soil with higher NH4+ availability favours AOB [66]. Despite a decrease in pH in the control (CK) treatment, AOB and AOA remained dominant. The accumulation of NH4+ in both neutral and alkaline soils due to BM biochar amendment appears to have suppressed the dominance of soil AOA and increased the abundance of soil AOB. This suggests that AOA is more dominant and active in NH4+-deficient soils, while AOB thrives in NH4+-rich soils. [69] reported similar findings, showing that the accumulation of N in soil decreases the abundance of soil AOA, while increasing the abundance of soil AOB. AOA is better suited to soil conditions characterized by acidity and nitrogen deficiency [70], where heterotrophic bacteria compete with nitrifying bacteria for soil inorganic N. Hence, the availability of N as an energy source promotes the AOB community. Our findings affirm that AOB will thrive in soils with higher NH4+ accumulation, further confirming the importance of N availability on the abundance of AOB [71]. Compared to other amendments, soils incubated with neem seed cake were dominated by Comammox bacteria abundance (Fig. 7). This was attributed to the acidity induced by neem seed cake following decomposition and organic N enrichment. In general, AOA typically dominates NH4+ oxidation in acidic soils [72]. However, recent studies have proven that Comammox bacteria outnumber AOA and can show nitrification activity in acidic soils [73, 74]. [75] also showed that Comammox bacteria are more abundant in acidic—N rich soil environments and have high tolerance to bioactive antibacterial compounds typically released by neem seed cake during decomposition, compared to AOA and AOB. The accumulation of inorganic nitrogen and further decrease in soil pH in the control treatment (CK) appear to have provided a conducive environment for Comammox to thrive in the acidic soil [76]. The abundance of Comammox was positively correlated with the mean nitrification rate and NO3−-N (Table S5). [76] reported similar findings, showing that the abundance of Comammox was positively correlated with soil N content. This implies that Comammox bacteria may directly compete with AOA and AOB for substrates, and its survival and growth could outperform other traditional ammonia-oxidizing bacteria [73].
Relationships between abundance of soil nitrifying microorganisms and soil chemical properties
Pearson correlation analysis results indicated that soil organic matter (SOM), NH4+, and NO3−N significantly influenced the abundance of AOA, AOB, and Comammox bacteria communities under various amendments across soil types (Table S5). [77] found similar results, showing that SOM was the primary driver of the AOA community, and AOA was more sensitive when the mineralization of residual organic matter released NH3 [78]. Furthermore, we discovered that NO3−-N had a more substantial impact on the AOB community abundance (Table S5). Previous studies have shown that soils with high N content support AOB proliferation [79]. [80] also suggested that NO3−-N is a crucial element affecting AOB diversity. The nitrification of NH4+ to NO3− provides a direct substrate for ammonia oxidizers, thereby stimulating the growth of ammonia oxidizers. This suggests that AOB rapidly oxidizes NH4+-N to NO3−-N [81, 82], generating protons and NO3−. Contrary to the findings of [83], our study found no significant relationship between soil total nitrogen and the abundance of Comammox bacteria. N availability has often been shown to significantly affect Comammox abundance [75], due to Comammox bacteria's higher metabolic capacity in soils with high N availability [84]. However, Comammox bacteria had a more significant relationship with soil MNR (Table S5). Soil MNR was an indicator of overall nitrification activity during incubation, and NH4+-N was a key factor affecting MNR [85]. Comammox bacteria has a higher affinity for NH4+ substrate than AOA and AOB [86], thrives in both acidic and alkaline soils, and uses NH4+ as the reaction substrate in the nitrification process, similar to AOB.
Conclusion
This study provides valuable insights into the complex interactions between soil amendments, soil properties, and the dynamics of N transformation processes. The application of BM-biochar and neem seed cake demonstrated contrasting effects on soil pH, N dynamics, and the abundance of nitrifying microbial communities across different soil types. In neutral and alkaline soils, the synergistic effects of BM-biochar and neem seed cake combination led to an accumulation of NH4+ and a temporary inhibition of nitrification, attributed to factors such as increased surface area and functional groups of BM-biochar, altered soil chemistry, and microbial competition for denitrification. Conversely, in acidic soil, this combination amplified and stimulated NO3− content, potentially due to enhanced heterotrophic nitrification and altered nitrifying microbial communities. The abundance of AOA, AOB, and Comammox bacteria was influenced by changes in soil pH and nitrogen availability induced by the amendments. AOB dominated in neutral and alkaline soils with higher NH4+ accumulation, while Comammox bacteria thrived in acidic soils amended with neem seed cake, likely due to their tolerance to bioactive compounds and ability to outcompete AOA and AOB under such conditions. Relationships between SOM, NH4+, NO3−, and the abundance of nitrifying microorganisms suggests that these soil properties play crucial roles in shaping the nitrifying microbial community. Furthermore, the co-amendment of BM-biochar and neem seed cake exhibited a quadratic increase in NH3 volatilization in neutral and alkaline soils, driven by the maintenance of favourable pH ranges and microbial activity. Overall, our study highlights the intricate interplay between soil amendments, physicochemical properties, and microbial communities in regulating N transformations and losses. Careful consideration of these factors is essential for developing effective soil management strategies that aim to optimize N retention and minimize environmental impacts.
Acknowledgements
This work was carried out with the support of Central Laboratory Facility at Adekunle Ajasin University Akungba-Akoko. Authors acknowledged the anonymous reviewers for comments to improve the quality of this work.
Author contribution
All authors contributed to the study conception and design. Manuscript preparation, data collection and analysis were performed by SO, AW, AA, and TA. The first draft of the manuscript was written by SO. ACA, AW, AA and TA edited and commented on previous versions of the manuscript. All authors read and approved the final manuscript.
Funding
This work did not receive any internal or external financial support.
Data availability
The datasets used or analyzed during the current study are available from the corresponding author on reasonable request.
Declarations
Competing interests
The authors declare no competing interests.
Publisher's Note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
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Abstract
In a 28-day incubation study, ball milling technologies were applied to enhance the sorptive and functional properties of pristine biochar. The effect of ball-milled (BM) biochar, neem seed cake, and their co-amendment was evaluated on nitrification, ammonia (NH3) volatilization, and the abundance of nitrifying microbial communities in three contrasting tropical soils of different pH (acidic, neutral, and alkaline). The amendments were applied at 2% dry w/w to soils fertilized with ammonium chloride (NH4Cl). Results showed that in neutral and alkaline soils, the co-amendment led to a 40% and 64% increase in NH3 volatilization, respectively, compared to control due to significant ammonium (NH4+) retention and temporary nitrification inhibition. Conversely, in acidic soil, BM biochar and neem seed cake amplified nitrification by 23% and 62%, respectively, compared to sole amendments, while neem seed cake increased NH3 volatilization by 56% compared to BM biochar + neem seed cake due to NH4+ retention, altered soil pH, and changes in nitrifying microbial community. The abundance of ammonia-oxidizing archaea (AOA), ammonia-oxidizing bacteria (AOB), and complete ammonia oxidizers (Comammox) was altered by changes in soil pH and N availability modulated by BM biochar and neem seed cake. Correlation analysis revealed significant relationships between soil organic matter (SOM), NH4+, and NO3− on the abundance of nitrifying microorganisms. The study affirms the efficacy of BM biochar-neem seed cake co-amendment on nitrification inhibition but indicates potential N losses by NH3 volatilization depending on soil type, highlighting the need for soil type-specific management strategies to optimize N retention while minimizing environmental impacts.
Article highlights
Ball-milled biochar + neem cake inhibited nitrification but increased NH3 volatilization in neutral and alkaline soils
In acidic soil, the co-amendment increased nitrification while reducing NH3 volatilization compared to neem cake alone
Effects on N cycling and microbial communities varied by soil pH, emphasizing need for soil-specific strategies
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Details
1 University of Lincoln, School of Natural Sciences, Lincoln, UK (GRID:grid.36511.30) (ISNI:0000 0004 0420 4262); University of Lincoln, Lincoln Institute for Agri-Food Technology, Lincoln, UK (GRID:grid.36511.30) (ISNI:0000 0004 0420 4262); Adekunle Ajasin University, Department of Agronomy, Faculty of Agriculture, Akungba Akoko, Nigeria (GRID:grid.442500.7) (ISNI:0000 0001 0591 1864)
2 University of Delaware, Department of Plant and Soil Sciences, Newark, USA (GRID:grid.33489.35) (ISNI:0000 0001 0454 4791)
3 Adekunle Ajasin University, Department of Agronomy, Faculty of Agriculture, Akungba Akoko, Nigeria (GRID:grid.442500.7) (ISNI:0000 0001 0591 1864)
4 Adekunle Ajasin University, Department of Forestry and Wildlife Management, Akungba Akoko, Nigeria (GRID:grid.442500.7) (ISNI:0000 0001 0591 1864)