Pathways and mechanisms for efficient nitrogen utilization in the high-temperature tolerant heterotrophic nitrifying bacterium Aeribacillus pallidus sp. GW-E

Abstract

This study isolated a novel thermophilic heterotrophic nitrifying bacterium, Aeribacillus pallidus sp. GW-E, from aerobic composting. Under conditions of 55 °C, the utilization efficiency of NH₄⁺-N, NO₃⁻-N, and NO₂⁻-N were 87.42%, 21.44%, and 51.68%, respectively. Whole-genome analysis identified key nitrogen metabolism genes (amt, npd, nirA, gdhA, glnA, and gltBD) as well as heat stress-related genes (GRPE, hslO, groES, groEL). Response surface optimization revealed that under conditions of a C/N ratio of 15, a temperature of 54 °C, and a pH of 8, the NH₄⁺-N utilization efficiency reached 100%. Enzyme activity assays indicated that the activities of three enzymes in the ammonia assimilation pathway were GS 1.014 ± 0.030 U/mg, GDH 1.114 ± 0.090 U/mg, and GOGAT 11.611 ± 0.061 U/mg, which were significantly higher than those of other pathways (P < 0.05). Nitrogen balance analysis confirmed that approximately 40.04% of the nitrogen was assimilated. In conclusion, the bacterium primarily utilizes ammonia assimilation, with additional assimilated nitrate reduction and nitrification pathways for nitrogen transformation. This strain represents a valuable microbial resource and provides a theoretical basis for nitrogen retention in high-temperature composting systems.

Supplementary Information

The online version contains supplementary material available at 10.1186/s12866-025-04415-4.

Introduction

With the advancement of global animal husbandry, the substantial accumulation of livestock manure and wastewater has led to serious environmental issues, including water, air, and soil pollution [1]. It is estimated that by 2030, the volume of livestock manure will reach 3.8 billion tons, containing a significant amount of organic matter (OM), nitrogen, phosphorus, and other nutrients that can be effectively utilized [2, 3]. However, the overall utilization rate remains below 60%, contributing to increased environmental pollution. Among the methods for the safe resource utilization of livestock manure, aerobic composting is prominent [4]. Nevertheless, traditional aerobic composting techniques frequently result in significant nitrogen loss due to inadequate nitrogen conservation strategies [5]. Studies have shown that the incorporation of heterotrophic nitrifying microorganisms effectively reduces nitrogen loss by lowering NH₄⁺-N levels, increasing NO₃⁻-N concentrations, and decreasing ammonia volatilization, thereby improving compost quality [6, 7]. Notably, ammonia emissions during the thermophilic phase can account for 21–77% of total nitrogen loss [8]. Autotrophic nitrifiers, due to their strict mesophilic nature, cannot adapt to the high temperatures (>50 °C) and localized anaerobic conditions present in aerobic composting, which limits their nitrogen transformation efficiency [9, 10] Similarly, mesophilic heterotrophic nitrifiers demonstrate reduced nitrification capacity under these conditions [11]. Therefore, the use of thermophilic heterotrophic nitrifying microorganisms is essential for minimizing nitrogen loss during the thermophilic phase.

Several thermophilic heterotrophic nitrifying bacteria have been identified, including Bacillus sp., Geobacillus sp., Bacillus methylotrophicus, and Aliibacillus thermotolerans strain BM62T [12–15]. However, these strains typically exhibit low utilization rates for NH₄⁺-N. For instance, Brevibacillus agri N2 and Gordonia paraffinivorans N52 demonstrate an average ammonium nitrogen utilization rate of less than 1 mg/L/h, even at a concentration of 99.64 mg/L [16, 17]. This rate is significantly lower than that of mesophilic heterotrophic nitrifying strains, such as Paracoccus denitrificans R-1 (9.94 mg/L/h) and Rhodococcus erythropolis strain Y10 (9.69 mg/L/h) [18, 19]. Studies have indicated that inoculating compost with 1% nitrifying bacteria (NTB) can reduce nitrogen loss by approximately 35% [20]. Additionally, ammonia-oxidizing bacteria (AOB) can reduce nitrogen loss from 53.83% to 47.08% [21]. However, mesophilic strains demonstrate very low survival rates during the high-temperature phase, which significantly limits their practical application. In aerobic composting, nitrogen loss primarily occurs during the high-temperature phase, mainly through NH₃ volatilization (46.10%–77.5%) [22], followed by the emission of greenhouse gases such as N₂ and N₂O. Experimental results indicate that the addition of thermophilic nitrifying bacteria can decrease nitrogen loss in compost by 29.7%–35.9%. However, their nitrogen retention efficiency is still constrained by the nitrogen utilization capacity of the strains themselves. This process is closely related to the intensity of nitrification-denitrification reactions driven by the compost microbial community [1, 7]. Therefore, the use of efficient thermophilic heterotrophic nitrifying bacteria to mitigate nitrogen loss during composting is scientifically well-supported, making the screening and application of such strains of paramount importance.

Current research on nitrogen transformation by heterotrophic nitrifying microorganisms has primarily focused on mesophilic bacteria, as well as strains that are tolerant to salt, cold, and heavy metals. These bacteria exhibit varying nitrification capabilities and contribute to the process by secreting specific enzymes [23]. The main nitrogen transformation pathways identified include nitrification, denitrification, ammonia assimilation, and dissimilatory nitrate/nitrite reduction [24]. Key enzymes involved in these processes are ammonia monooxygenase (AMO), hydroxylamine oxidoreductase (HAO), nitrite reductase (NIR), and nitrate reductase (NAR). The classic nitrification pathway involves the conversion of NH₄⁺-N to NH₂OH and NO₂⁻-N, followed by further oxidation to NO₃⁻-N. Denitrification proceeds through the following steps: NO₃⁻ → NO₂⁻ → NO → N₂O → N₂, with N₂O ultimately being converted to N₂ [24, 25]. In dissimilatory nitrate reduction, NO₃⁻ was reduced to NO₂⁻, and NO₂⁻-N was converted to NH₄⁺-N, which was then transformed into gases like N₂ and N₂O. In assimilatory nitrate reduction, NH₄⁺-N supports cell growth through the action of glutamate dehydrogenase (GDH) [26]. Heterotrophic nitrifying bacteria typically assimilate 41.7–67.8% of NH₄⁺-N into organic nitrogen [27, 28], with relevant genes including glnA, gdhA, gltB, and gltBD [29]. Elevated temperatures disrupt microbial metabolism by affecting enzyme activity, gene expression, and cellular structure [30]. This indicates that thermal stress may alter the nitrogen transformation pathways in thermophilic heterotrophic nitrifying bacteria. However, research on these strains is currently insufficient, leading to limited nitrogen utilization efficiency, which hinders their application in aerobic composting and biological nitrogen-retention inoculants. Therefore, elucidating the nitrogen transformation mechanisms of thermophilic heterotrophic nitrifying bacteria is of significant importance.

In this research, we extracted a thermophilic, heterotrophic nitrifying bacterium from the elevated temperature stage of aerobic composting. The goals of this research were to: (1) To investigate the nitrogen transformation capabilities of the isolated strain under high-temperature conditions using NH₄⁺-N, NO₃⁻-N, and NO₂⁻-N as sole nitrogen sources; (2) Explain the nitrogen transformation processes and heat resistance strategies of the strain in high-temperature environments by examining enzyme activity, conducting nitrogen balance experiments, and performing whole-genome analysis. (3) To explore the effects of different carbon sources, temperature, C/N ratio, and pH conditions on the nitrogen transformation performance of the strain, and to further optimize the cultivation conditions of the strain using response surface methodology. This study provides new resources for high-temperature heterotrophic nitrifying bacteria and offers in-depth insights into nitrogen transformation pathways and mechanisms, thereby providing scientific evidence for the application of this strain.

Materials and methods

Isolation and screening

This study utilized an enriched medium (DM) to isolate nitrifying heterotrophic bacteria. The medium composition was as follows: 2 g of ammonium sulfate, 14.31 g of sodium succinate, and 50 mL of Vischer’s salt solution. A 10 g sample of aerobic compost from a farmer cooperative specializing in cattle and sheep breeding in Guanghe County, Linxia Hui Autonomous Prefecture, Gansu Province, was inoculated into 200 mL of LB medium (containing 10 g of peptone, 5 g of yeast extract, and 10 g of sodium chloride) and incubated at 55 °C with agitation at 160 rpm for 5 days. This process was repeated for three successive transfers. After the final transfer, 2 mL of the enriched culture was transferred to the DM medium, and three additional serial transfer were carried out under the same conditions. The culture was then serially diluted (10⁻³ to 10⁻⁵) and spread onto solid DM agar plates (DM supplemented with 18 g/L agar), followed by incubation at 55 °C for 3–5 days. Colonies of distinct mophologies were repeatedly streaked to obtain pure isolates by single colonies. The purified isolates were re-inoculated into the enrichment medium, and 1 mL of culture subjected to the Nessler’s reagent colorimetric assay on a porcelain plate to preliminarily screen for strong decolorization (indicative of NH₄⁺-N removal). Candidate strains were further evaluated for growth rate and NH₄⁺-N utilization efficiency and the best-performing isolate was selected. Nitrogen balance analysis was subsequently conducted to determine ammonia assimilation, and the strain exhibiting the highest assimilation rate was designated GW-E for downstream studies. All experiments were performed in triplicate. The final strain was stored in glycerol solution at −80 °C for long-term preservation.

Strain identification

Bacterial DNA was obtained using a DNA extraction kit (D1600; Solely Bio., Beijing, China). The identification in molecular biology was conducted following the methods described by Zhao [16]. Scanning electron microscopy (SEM) was performed at Wuhan Saiwei Biotech Co., Ltd. The PCR products were examined through 0.5% agarose gel electrophoresis, and the amplified products were sequenced by Shanghai Shenggong Bioengineering Co., Ltd. The sequencing outcomes were analyzed for similarity using the Basic Local Alignment Search Tool (BLAST) with the NCBI database (http://blast.ncbi.nlm.nih.gov/Blast.cgi), and the sequences were also uploaded to the GenBank database. Ultimately, a phylogenetic tree was created utilizing the neighbor-joining approach in MEGA 11.0 software.

Study on the conversion performance of strains on different nitrogen sources

The initial nitrogen concentration—supplied as NH₄⁺-N, NO₃⁻-N, or NO₂⁻-N—was 400 mg N·L⁻¹ in each of the three media (HNM, ADM-1, and ADM-2). Sodium succinate served as the sole carbon source, and the carbon-to-nitrogen (C/N) ratio was fixed at 10. Prior to the experiment, 2 mL (1% v/v) of seed culture was inoculated into 250 mL Erlenmeyer flasks containing 200 mL of DM medium and incubated at 55 °C and 180 rpm for 48 h, until the optical density at 600 nm (OD₆₀₀) reached approximately 1.0. A 10 mL aliquot of the culture was centrifuged at 10,000 rpm for 5 min at 4 °C, and the cells were collected and washed three times with sterile water and phosphate-buffered saline (PBS). The cells were then resuspended to an OD₆₀₀ of approximately 1.0. The washed cell suspension was inoculated at a 1% inoculum rate (2 mL) into three different test media (HNM, ADM-1, and ADM-2, each 200 mL in 250 mL Erlenmeyer flasks) and incubated at 55 °C with shaking at 180 rpm for 96 h. Samples were taken at 0, 24, 48, 72, and 96 h to measure OD₆₀₀, ammonium nitrogen (NH₄⁺-N), nitrate nitrogen (NO₃⁻-N), nitrite nitrogen (NO₂⁻-N), dry cell weight (DCW), and cellular nitrogen content. OD₆₀₀, NH₄⁺-N, NO₃⁻-N, and NO₂⁻-N were measured using spectrophotometric methods, sodium salicylate reagent, ultraviolet spectrophotometry, and N-(1-naphthyl)ethylenediamine spectrophotometry, respectively [31]. DCW was determined by centrifuging and washing the cells, followed by drying at 80 °C to a constant weight [32]. Cellular nitrogen content was measured using the Kjeldahl method [33]. All experiments were conducted under sterile conditions, with un-inoculated medium serving as the blank control, and each experiment was performed in triplicate (Table 1).

Table 1.

Content of different nitrogen source culture media

Medium	Composition and content (g)				Trace element	Distilled water	Concentration(mg/L)
	C4H4Na2O4	(NH4)2SO4	KNO3	NaNO2
Heterotrophic nitrification medium(HNM)	20.18 g	1.89 g	-	-	50 mL	950 mL	416.31
Aerobic denitrification medium(ADM-1)	20.18 g	-	2.88 g	-	50 mL	950 mL	409.68
Aerobic denitrification medium(ADM-2)	20.18 g	-	-	1.96 g	50 mL	950 mL	415.71
Enriched domestication medium(DM)	14.31 g	2 g	-	-	50 mL	950 mL	424.00

Trace element(g/L) :K₂HPO₄ 5, MgSO₄·7H₂O 2.50, NaCl 2.50, FeSO₄·7H₂O 0.05, MnSO₄·4H₂O 0.05。

Nitrogen balance experiment

To investigate the nitrogen metabolism characteristics of this strain, nitrogen balance experiments were conducted using NH₄⁺-N, NO₃⁻-N, and NO₂⁻-N as the sole nitrogen sources. Following the method outlined in Sect. 2.3, 2mL of the bacterial suspension with an optical density (OD) at 600 nm of approximately 1.0, was inoculated into HNM, ADM-1, and ADM-2 media and incubated at 55 °C while shaking at 160 rpm for 96 h. Total nitrogen (TN) and total dissolved nitrogen (TDN) in the supernatant at 0 h and 96 h were determined using the alkaline potassium persulfate digestion method [34]. TDN measurements required filtration of the bacterial suspension through a 0.22 μm sterile membrane to remove the cells. The nitrogen balance data were calculated using the subtraction method, where total nitrogen in the supernatant (TDN), cellular nitrogen (Cell-N) and Gaseous-N were determined. Based on these values, the assimilation ratio and gas ratio of the strain were estimated. Assimilation efficiency and denitrification efficiency were then derived, with the calculation method detailed in Sect. 2.8.2. All experiments were conducted under sterile conditions, with un-inoculated medium serving as the blank control, and each treatment was repeated three times.

Enzyme activity assay

The bacterial suspension that had been pre-cultured was added to the enrichment medium, and the GW-E strain was incubated at 55 °C for 48 h. The bacterial suspension was then centrifuged at 10,000 rpm for 20 min at a temperature of 4 °C and washed three times with 0.01 mol/L PBS (pH 7.4). The suspension was then resuspended and subjected to ultrasonic cell disruption to obtain a cell-free extract. The extract was re-centrifuged at 10,000 rpm for 20 min at 4 °C, and the supernatant was collected as the crude enzyme solution. The activities of AMO, HAO, NIR, NAR, NOR, NOS, glutamine synthetase (GS), glutamate synthase (GOGAT), and GDH were measured using bacterial ELISA kits (ADS-W-N002-48; Jingmei, Jiangsu, China). Each experiment was conducted in triplicate.

Whole-Genome sequencing and functional annotation

Bacterial genome sequencing was performed by Shanghai Meiji Biotechnology Co., Ltd. using PacBio Sequel II and Illumina platforms for bioinformatics data analysis. The entire genome of the strain GW-E was sequenced, with a total length of 3,821,580 bp (Supplementary Table S5). The coding sequences (CDSs) within the genome were predicted using Glimmer (v3.02), and the plasmid genes were predicted using GeneMarkS software. tRNAs were predicted using tRNAscan-SE (v2.0), and rRNAs were predicted using Barrnap (https://github.com/tseemann/barrnap). Functional annotation of the predicted CDSs was carried out using sequence alignment tools, such as BLASTP, Diamond, and HMMER, and functional information was obtained from the NR, GO, COG, and KEGG databases. A circular genome map of GW-E was constructed using Circos (version 0.69–6.69). All analyses were conducted using the I-Sanger cloud platform (www.i-sanger.com) provided by Shanghai Meiji Biotechnology.

Optimization of culture conditions for the strain

Single-Factor experiment

To evaluate the impact of different environmental conditions on the nitrogen utilization capacity of the strain, a series of experiments were conducted under a nitrogen content of 400 mg/L, with the carbon source, carbon-to-nitrogen ratio (C/N), pH, and temperature as variables. Glucose, sodium citrate, sodium succinate, sucrose, and sodium acetate were used as carbon sources to investigate their effects on the strain. The C/N ratio was adjusted from 5:1 to 25:1 to investigate how it influences the nitrogen utilization capacity of the strain. The pH was adjusted to 5, 6, 7, 8, 9, and 10 to observe its influence on the nitrogen utilization capacity of the strain. All experiments were conducted at 55 °C and 160 rpm for 96 h. To explore the impact of temperature on the strain’s nitrogen utilization capacity, experimental temperatures were set at 45 °C, 50 °C, 55 °C, 60 °C, and 65 °C, with the culture maintained at 160 rpm for 96 h. For all these experiments, samples were taken every 12 h to measure OD₆₀₀, NH₄⁺-N, NO₃⁻-N, and NO₂⁻-N, with the same measurement methods as described in Sect. 2.3. All experiments were performed under sterile conditions, with three replicates for each trial, and an uninoculated culture medium was used as a control.

Response surface optimization

Based on a single-factor experiment, temperature, pH, and the carbon-to-nitrogen (C/N) ratio were identified as the factors with the greatest impact on strain, and the appropriate range for each factor was determined. The NH₄⁺-N utilization efficiency was designated as the final response variable, and the variation in NH₄⁺-N utilization efficiency was analyzed in relation to the three identified factors. A Box-Behnken design featuring three factors and three levels was employed to optimize the culture conditions for the strain. The experiment consisted of 17 trials (see Supplementary Table S4). The NH₄⁺-N utilization efficiency (%) served as the response variable to evaluate the effects of the three factors—temperature (A), pH (B), and C/N ratio (C)—on the response value and to investigate their interactions, with the aim of determining the optimal culture conditions (Table 2).

Table 2.

Each factor in the response surface molecule

Factor		level
	−1	0	1
A-temperature	50℃	55℃	60℃
B-pH	7	8	9
C-C/N	5	10	15

Data analysis and calculation methods

Data analysis

The data were analyzed statistically with Microsoft Excel, and a one-way analysis of variance (ANOVA) was conducted using IBM SPSS Statistics 26.0. The results are presented as the mean ± standard deviation. Statistical significance was considered at P < 0.05, while a P-value greater than 0.05 indicated no significant difference. Data analysis and graph creation were conducted using Origin, version 2024. Pearson correlation analysis was conducted using IBM SPSS Statistics 26.0, yielding correlation coefficients that varied from − 1 to + 1. The closer the value is to ± 1, the stronger the correlation. Design-Expert software (version 11.0) was used for the Box-Behnken Design(BBD) response surface.

Calculation methods

The calculation formulas for NH₄⁺-N, NO₃⁻-N, and NO₂⁻-N, N utilization efficiency and N utilization rate are as follows [16]:

• N utilization efficiency (%) = (C_{0 h}−C_{t h})/C_{0 h}×100%;

• N utilization rate (mg/L·h) = (C_{0 h}−C_{t h})/t.

• Cumulative production of NO₃⁻-N, NO₂⁻-N (mg/L)=(C_t – C₀),

• Where C_{0 h}, C_{t h} are the initial and final concentrations of each nitrogen compound.

• (e.g., NH₄⁺-N, NO₃⁻-N, NO₂⁻-N) at 0 h and t h, respectively.

The formula for calculating nitrogen balance was obtained from Zhang [35]. The calculation formulas are as follows:

• Cell nitrogen 0 h (mg/L) = TN_0h − DTN_{0 h}.

• Cell nitrogen 96 h (mg/L) = TN_{96 h} − DTN_{96 h}.

• Cell nitrogen (mg/L) = Cell nitrogen _96 h − Cell nitrogen _0 h;

• Gaseous-N(mg/L) = TN _0 h- TN_96h;

The calculation formula is as follows, and the assimilation efficiency and aenitrification efficiency of N transformation by the strain is calculated by the following formula:

• Assimilation efficiency (%) = Cell nitrogen/DTN_{0 h} × 100%

• Aenitrification efficiency (%) = Gaseous-N/DTN_{0 h} × 100%

Results and discussion

Isolation and identification of the strain

The strain GW-E appeared white on the isolation medium, exhibiting slightly raised colonies with regular edges; Gram staining was negative. Figure 1a and b display the isolation and SEM images of strain GW-E. The 16 S rRNA gene sequence obtained was submitted to NCBI, and a homology search was conducted using BLAST. The 16 S rRNA gene sequence of strain GW-E exhibited 100% similarity to that of Aeribacillus pallidus strain MRP112 (GenBank accession number PQ448334). A phylogenetic tree was created using MEGA 11.0 based on the 16 S rRNA gene sequence (Fig. 1c), which further validates the classification of the strain as Aeribacillus pallidus sp. GW-E.

Fig. 1.

Isolation map of strain GW-E, scanning electron microscopy image, and 16S rRNA gene phylogenetic tree. Note:(a) Plate culture image of strain GW-E; (b) Scanning electron microscopy image; (c) Phylogenetic tree constructed from the 16S rRNA gene sequence of strain GW-E. The numbers in parentheses are GenBank accession numbers

Analysis of nitrogen source utilization characteristics of the strain GW-E

Utilization of ammonia nitrogen as the sole nitrogen source

Figure 2(a) illustrates the growth curve and nitrogen transformation of strain GW-E when NH₄⁺-N served as the sole nitrogen source. After 72 h, the OD₆₀₀ value peaked at 1.42, which was significantly higher than the values recorded at other time intervals (P < 0.05). Furthermore, the cell dry weight (DCW) exhibited a notable increase between 72 and 96 h. Throughout the study period, the NH₄⁺-N concentration decreased, with an NH₄⁺-N utilization efficiency of 87.42% and an average utilization rates of 3.79 mg/L/h, which surpassed the rates reported for Bacillus simplex h-b (0.74 mg/L/h) [36] and Klebsiella sp. TSH15 (2.37 mg/L/h) [37]. The accumulation of NO₃^--N and NO₂^--N was widely recognized as a crucial indicator for assessing the occurrence of nitrification [38]. Significant accumulations of NO₃^--N (84.32 ± 0.31 mg/L at 72 h) and NO₂⁻-N (30.13 ± 0.44 mg/L at 96 h) were detected in the culture system, with their accumulation levels significantly exceeding the reported values for the known heterotrophic nitrifiers Sneathiella aquimaris 216LB-ZA1-12T and Pseudomonas mendocina X49 [39, 40]. As illustrated in Fig. 2(d). A significant negative correlation (P < 0.01) was observed between NH₄⁺-N concentration and the concentrations of NO₃^--N and NO₂^--N, indicating that the strain GW-E exhibited a pathway for the utilization of NH₄⁺-N that can be converted into NO₃^--N and NO₂^--N [41]. As illustrated in Figs. 3 and 193.18 ± 1.13 mg/L of nitrogen was assimilated into cellular biomass during the NH₄⁺-N transformation process, representing 48.29% of the total nitrogen. A dynamic change was also observed, with the concentration of NO₂^--N decreasing from 24.56 mg/L to 12.65 mg/L. Furthermore, a significant negative correlation (P < 0.01) was found between NO₂^--N levels and both OD₆₀₀ and DCW, indicating that the strain may assimilate NO₂^--N into biological nitrogen for cell growth through metabolic pathways similar to those of Bacillus cereus GS-5 and Halomonas venusta SND-01 [42]. In summary, the strain GW-E exhibited a dual NH₄⁺-N metabolic pathway: on one hand, it assimilated ammonium nitrogen into cellular biomass; on the other hand, the strain GW-E can convert ammonia nitrogen into nitrate and nitrite through the process of nitrification.

Fig. 2.

Growth pattern and nitrification profile of Aeribacillus pallidus sp. GW-E in the sole nitrogen source medium at 55◦C.NH₄⁺-N as the sole nitrogen source (a); NO₃⁻-N as the sole nitrogen source (b); NO₂⁻-N as the sole nitrogen source (c); stands for correlation analysis using NH₄⁺-N, NO₃⁻-N, and NO₂⁻-N as the sole nitrogen sources(d, e, f)

Fig. 3.

Nitrogen Content, Total Nitrogen Content, and Assimilation Efficiency under Different Nitrogen Source Utilization (a) Nitrogen Content, Total Nitrogen Content of Cell Dry Weight (b) Assimilation Efficiency under Different Nitrogen Source Utilization. Note: All experimental test datawere replicated three times and data were marked by mean ± standard error(SE). Cell dry weight total nitrogen (mg/L)= (Cell nitrogen(%) * DCW) ÷ 1000; Assimilation Efficiency(%) =(Cell dry weight total nitrogen ÷ 400)x 100

Utilization of nitrate nitrogen as the sole nitrogen source

The growth and nitrogen conversion profiles utilizing NO₃^--N as the sole nitrogen source are illustrated in Fig. 2(b). The OD₆₀₀ reached 0.73 ± 0.001 (P < 0.05) at 48 h, while the DCW showed the most significant biomass accumulation between 72 and 96 h, corresponding to the logarithmic growth phase of the strain GW-E. During this period, strain GW-E demonstrated the highest utilization efficiency of NO₃^--N at 21.44%, with a maximum utilization rates of 1.93 mg/L/h. This rate was notably higher than the 0.80 mg/L/h utilization rates of NO₃^--N observed in Janthinobacterium sp. J1-1 during its logarithmic phase [43]. After 48 h, the concentrations of NH₄⁺-N and NO₂^--N increased to 18.23 ± 0.29 mg/L (P < 0.05) and 431.33 ± 0.00 mg/L (P < 0.05), respectively, as illustrated in Fig. 2(e). A significant negative correlation (P < 0.05) was observed between the concentrations of NO₃^--N and NO₂^--N. During the logarithmic phase, strain GW-E exhibited an NH₄⁺-N concentration of 18.23 ± 0.29 mg/L, indicating that strain GW-E may be capable of converting NO₃^--N to NH₄⁺-N. This process is consistent with a nitrogen metabolic pathway that involves the dissimilatory reduction of nitrate to NH₄⁺-N [37, 44]. The increase in NO₂^--N concentration was hypothesized to result from the strain GW-E utilizing NO₃^--N as an electron acceptor, leading to nitrate reduction and its conversion to NO₂^--N. When the NO₂^--N concentration becomes excessively high, it is reduced to NH₄⁺-N through either assimilatory nitrate reduction to ammonium (ANRA) or dissimilatory nitrate reduction to ammonium (DNRA) [45]. Both processes contribute to an increase in NH₄⁺-N levels. A significant positive correlation (P < 0.05) was observed between NH₄⁺-N and NO₂^--N concentrations, indicating a pathway in which NO₂^--N is converted to NH₄⁺-N.

Utilization of nitrite nitrogen as the sole nitrogen source

When NO₂⁻-N was used as the sole nitrogen source, as shown in Fig. 2(c), the strain GW-E exhibited a significant growth lag, likely due to the cytotoxic effects of nitrite, which required a longer adaptation period for the microorganisms to acclimate to their environment [46]. Nevertheless, the strain demonstrated a notable capacity for nitrite metabolism. The utilization efficiency of NO₂⁻-N by strain GW-E was 51.68%, with an average utilization rates of 4.31 mg/L/h. This rate was significantly higher than the average utilization rates of 0.73 mg/L/h observed in Bacillus strain N31 after 42 h [47] and 0.76 mg/L/h in Acinetobacter sp. YT03 [48]. Notably, after 96 h of cultivation, NO₃⁻-N accumulation was detected at 338.01 ± 0.94 mg/L (P < 0.05), and a significant negative correlation was observed between the concentrations of NO₂⁻-N and NO₃⁻-N (P < 0.05, Fig. 2f). This finding indicated that the strain possesses the capability for nitrification, converting NO₂⁻-N to NO₃⁻-N. Additionally, NH₄⁺-N accumulation was detected at 72 h, measuring 12.79 ± 0.14 mg/L (P < 0.05). Considering that 34.46% of the nitrogen was assimilated into biomass nitrogen (Fig. 3b), these results suggest that the strain GW-E utilizes NO₂⁻-N through a nitrification pathway to convert it to NO₃⁻-N. Furthermore, through the assimilatory nitrate reduction pathway, NO₃⁻-N is reduced to NH₄⁺-N, which is subsequently assimilated into biological nitrogen via ammonium assimilation.

Nitrogen balance analysis

Previous studies have indicated that bacterial assimilation and nitrification processes do not lead to nitrogen loss within the culture system [49], suggesting that nitrogen loss primarily results from gas production. The nitrogen balance analysis (Table 3) demonstrated that the strain GW-E does not cause significant nitrogen loss during its nitrogen metabolism process. In the NH₄⁺-N medium, 40.04% of the nitrogen was assimilated into intracellular nitrogen, with an assimilation efficiency comparable to that of the halophilic strain Halomonas venusta SND-01 [42] and Candida boidinii L21 [50]. Notably, when NO₃⁻-N and NO₂⁻-N were used as nitrogen sources, 9.63% and 31.49% of total nitrogen (TN) were converted into cellular nitrogen for assimilation, This result differs from the findings for Glutamicibacter arilaitensis EM-H8 [51], which showed that around 77.88% of nitrogen was released as gas, while only 14.47% was transformed into biomass. Indicating that the primary assimilation pathway of strain GW-E utilizes nitrogen sources efficiently.

Table 3.

Nitrogen balance of strain GW-E in removing ammonia nitrogen, nitrate nitrogen, and nitrite nitrogen

Nitrogen source	Initial nitrogen(mg/L)			TN(mg/L)		DTN(mg/L)
	NH4+-N	NO2−-N	NO3−-N	Initial	Final	Initial		Final	Intracellular-N(mg/L)	Proportion (%)	Gaseous-N(mg/L)	Proportion(%)
NH4+-N	416.31 ± 0.00	7.22 ± 0.29	10.84 ± 1.15	440.27 ± 6.01	365.09 ± 23.30	433.26 ± 11.26		198.63 ± 29.85	173.47 ± 2.65	40.04%	75.19 ± 5.18	17.35%
NO3−−-N	15.61.±0.28	7.03 ± 0.49	437.27 ± 0.98	432.03 ± 51.75	296.63 ± 6.77	389.14 ± 100.12		302.06 ± 17.93	37.46 ± 6.16	9.63%	135.40 ± 0.02	34.69%
NO2−-N	17.58 ± 0.14	428.31 ± 0.14	35.60 ± 1.07	423.64 ± 2.52	320.55 ± 35.23	388.73 ± 18.50		221.24 ± 27.33	122.4 ± 42.24	31.49%	91.27 ± 10.93	23.48%

All experimental test datawere replicated three times and data were marked by mean ± standard error(SE). Intracellular nitrogen=(TN -DTN) _Final + (TN - DTN) _Initial; Proportion (%) = (Intracellular nitrogen ÷ DTN_Initial) x 100. Gaseous-N = TN _Initial- TN_Final; Proportion (%) = (Gaseous-N ÷ DTN_Initial) x 100

Enzyme activity analysis

To further investigate the nitrogen transformation mechanisms of strain GW-E, we systematically quantified key functional enzymes involved in the nitrification, denitrification, and ammonia assimilation pathways. Enzyme activity measurements (Fig. 4) revealed that strain GW-E exhibited specific activities of ammonia monooxygenase (AMO) and hydroxylamine oxidoreductase (HAO) of 0.32 ± 0.11 U/mg and 0.04 ± 0.01 U/mg, respectively. These values were significantly higher than those reported in the literature (AMO: 0.155 ± 0.009 U/mg; HAO: 0.037 ± 0.004 U/mg) [42, 52]. The specific activities of enzymes involved in the denitrification pathway were generally low, with nitrate reductase (NAR) at 0.05 ± 0.00 U/mg, nitrite reductase (NIR) at 0.04 ± 0.00 U/mg, and both nitric oxide synthase (NOS) and nitric oxide reductase (NOR) at 0.002 U/mg. This finding aligns with the typical characteristics of heterotrophic nitrifying bacteria, where nitrification is accompanied by low levels of denitrification [53]. This suggests that strain GW-E can efficiently utilize NH₄⁺-N.

Fig. 4.

Key enzyme activity of high-temperature resistant strain Aeribacillus pallidus sp.GW-E

The specific activities of key enzymes in the ammonium assimilation pathway—glutamine synthetase (GS), glutamate dehydrogenase (GDH), and glutamate synthase (GOGAT)—were significantly higher than those of other enzymes involved in nitrogen metabolism, measuring 1.014 ± 0.030 U/mg, 1.114 ± 0.090 U/mg, and 1.611 ± 0.061 U/mg, respectively. These results support the previous hypothesis that strain GW-E possesses an effective ammonium assimilation process, with the highest activity observed for GOGAT. This phenomenon may be linked to the strain’s unique metabolic regulatory mechanism: under high-temperature stress conditions, the strain preferentially utilizes small carbon sources, such as sodium fumarate, to enter the tricarboxylic acid cycle, thereby promoting the production of α-ketoglutarate and generating a substantial amount of energy [54]. To maintain the energy balance necessary for normal growth, the strain significantly upregulates the activity of GS and GOGAT through a feedback regulatory mechanism. Furthermore, although GS typically exhibits a higher affinity for ammonia than GDH, GDH plays a more prominent role in ammonia assimilation under high ammonia conditions [55, 56], which aligns with the observation that GDH activity was slightly higher than that of GS in this study.

In summary, the strain GW-E demonstrated a high capacity for nitrification and efficient ammonia assimilation. Its unique enzymatic activity suggests that this strain can effectively reduce nitrogen volatilization during aerobic composting, underscoring its significant potential for practical applications.

Whole genomic sequencing and molecular mechanism of nitrogen transformation

Molecular mechanism analysis of nitrogen metabolic pathway in strain GW-E

The whole genome sequencing results revealed that the genome size of strain GW-E is 3,821,580 bp, with a GC content of 39.08% and a total gene length of 3,113,958 bp (Supplementary Table S5). In terms of genome annotation, 2,990 protein-coding genes were successfully identified through comparison with the COG database (Supplementary Table S8). Among these, 11 genes were involved in nitrogen metabolism, and 23 genes were related to temperature adaptation. The nitrogen metabolism-related genes include amt (ammonium transporter), ndhF (core subunit 5 of ubiquinone oxidoreductase), npd (nitrate monooxygenase), nirA (ferredoxin-dependent nitrite reductase), among others, as well as genes involved in ammonia assimilation pathways, such as glnA, gudB, gdhA, gltB, and gltD. Based on the comprehensive genomic information and annotation results, it can be inferred that strain GW-E utilizes NH₄⁺-N primarily through three pathways: (1) the nitrification pathway, (2) the ammonia assimilation pathway, and (3) the assimilatory nitrate reduction pathway. A schematic diagram of the nitrogen transformation pathways is presented in Figs. 5, 6 and 7.

Fig. 5.

Circular map of the strainGW- E genome. From the outer circle to the inner circle: (1) the size of genome; (2–3): the coding sequences on the positive and negative strands, respectively (different colors mean different COG functional categories); (4) tRNA (orange) and rRNA (blue); (5) G + C content (red above average, blue below average); (6) GC skew (green means G%>C%, orange means C%>G%)

Fig. 6.

A map of the nitrogen metabolism pathways in strainE annotated using the KEGG database. Red-bordered boxes indicate the annotated genes, and thenumbers within some of these boxes are the annotated enzyme commission numbers

Fig. 7.

Schematic diagram of nitrogen metabolism pathway of the strain GW-E

The first pathway is the ammonia assimilation pathway. We successfully annotated all the key genes involved in this pathway, including Glutamate Dehydrogenase (GDH), Glutamine Synthetase (GS), and Glutamine Oxoglutarate Aminotransferase (GOGAT). These genes play a crucial role in regulating the ammonia assimilation process, thereby controlling the intracellular nitrogen pool [57]. The specific transformation process is as follows: First, the ammonium transporter protein (AMT), encoded by the amt gene, is responsible for transporting a significant amount of NH₄⁺-N across the cell membrane, thereby initiating the ammonia assimilation process. Next, the first reaction of glutamine synthetase (GS), catalyzed by the glnA gene, combines ammonia with glutamate to form glutamine, effectively converting inorganic nitrogen into organic nitrogen. Subsequently, the enzyme glutamate synthase (GOGAT), catalyzed by the glnB gene, mediates an amino transfer reaction that converts glutamine into two molecules of glutamate. These two molecules of glutamate can then serve as substrates for GS, providing energy for bacterial growth and metabolism while maintaining the normal growth and metabolic activities of the bacteria [56–58].

The second pathway is the assimilatory nitrate reduction pathway. The utilization of nitrite by strain GW-E primarily depends on the nirA gene, which encodes ferredoxin-dependent nitrite reductase (Fd-NiR). This enzyme catalyzes the conversion of NO₂⁻-N to NH₄⁺-N, facilitating the transformation of nitrite into biological nitrogen, which is then directly utilized by the strain. This process constitutes the assimilatory nitrate reduction pathway [59]. Studies have shown that the nirA gene is present in the strain Halomonas venusta SND-01, and its deletion results in the inhibition of nitrogen metabolic pathways. When nitrite nitrogen was used as the sole nitrogen source, only about 14.47% of nitrogen was converted into biological nitrogen, suggesting that strain GW-E also possesses the assimilatory nitrate reduction pathway [42, 60]. Given the critical role of the nirA gene in this pathway, future integration of gene knockout, gene recombination, and heterologous expression technologies may further enhance the nitrogen metabolic capacity of the strain.

The third pathway is the nitrification pathway. In the nitrogen source utilization tests of strain GW-E, it demonstrated good adaptability to the use of NH₄⁺-N, while its utilization rates for NO₃⁻-N and NO₂⁻-N were relatively low. Additionally, a significant conversion relationship was detected among NO₃⁻-N, NO₂⁻-N, and NH₄⁺-N during the utilization of NH₄⁺-N. However, genome analysis results revealed that strain GW-E does not annotate the key genes involved in nitrification, specifically amo and hao. A similar situation was observed in the genome analyses of Pseudomonas aeruginosa G16, Halomonas, Alcaligenes amioxyans HO-1, and Pseudomonas citronellolis YN-21, where these two genes were absent [61–64]. In Pseudomonas aeruginosa G16, it was found that a protein encoded by GE002604 possesses a 2 F-2 S conformation and a redox enzyme iron-sulfur cluster binding domain, which can functionally replace the AMO and HAO genes [61]. Similarly, gene 1608 and gene 1525 in the genome of strain GW-E also contain similar domains, suggesting the possible presence of unknown genes that could substitute for the typical functions of the nitrification pathway. Moreover, enzyme activity assays (Fig. 4) indicated that the nitrification enzyme activity in strain GW-E was higher than that of the denitrification process, further supporting the existence of a nitrification pathway. To address the absence of the amo and hao genes, future studies could combine transcriptomics and mass spectrometry to identify alternative genes involved in the ammonia oxidation pathway.

Molecular mechanism of thermotolerance in strain GW-E

The high-temperature adaptation mechanism of the strain GW-E involves a multi-layered molecular regulatory network. Genomic analysis has revealed that it possesses a comprehensive stress response system (see supplementary Table S7). To maintain protein homeostasis, a molecular chaperone network that includes HSP70 (GRPE), HSP33 (hslO), HSP10 (groES), and HSP60 (groEL) prevents protein aggregation and promotes refolding through an ATP-dependent mechanism [65, 66]. Simultaneously, the FtsH membrane protease and Clp family proteases (ClpB/ClpC) form a proteolytic system, establishing a “recognition-repair-degradation” tri-level defense system [67, 68]. Regarding genomic stability, DNA gyrase B (encoded by gyrB) repairs heat-induced damage, while the rbfA and punA genes are involved in transcriptional regulation, working in conjunction with adjustments in membrane lipid composition to maintain cellular integrity [69]. The ATP-dependent nature of heat shock proteins (HSPs) is linked to energy metabolism, and these proteins may protect key nitrogen metabolism enzymes, while the proteolytic system provides precursors for nitrogen assimilation [70]. Through these mechanisms, the strain GW-E maintains stable survival and continues to perform nitrogen conversion capabilities at 55 °C. However, the synergistic relationship between HSPs and nitrogen metabolism enzymes still requires further validation through transcriptomic analysis and protein-protein interaction studies.

Optimization of cultivation conditions for strain GW-E

Different cultivation conditions affecting the nitrogen transformation ability of strain GW-E

As shown in Fig. 8 (a), when sodium succinate was used as the carbon source, the optical density (OD₆₀₀) at the conclusion of the cultivation was measured at 1.26 ± 0.01 (P < 0.05), and the NH₄⁺-N utilization efficiency was 90.02%. In contrast, The highest NO₂^–-N concentration at 84 h was 69.06 ± 12.84 mg/L (P < 0.05). This indicated that the strain GW-E exhibits superior growth and utilization when sodium succinate was employed as the carbon source. Sodium succinate, being a small-molecule carbon source with a simple structure, can directly participate in the tricarboxylic acid (TCA) cycle, making it readily utilized by microorganisms. This enhances the microorganism’s ability to utilize nitrogen and promotes ammonia assimilation, ultimately converting it into organic nitrogen [71, 72].

Fig. 8.

Growth, ammonia nitrogen utilization efficiency, and cumulative production of nitrate and nitrite nitrogen by Aeribacillus pallidus sp. GW-E under different conditions (a) carbon sources; (b) C/N ratios; (c) pH; and (d) Temperature. Letters above columns indicate a significant difference at P < 0.05

As shown in Fig. 8 (b), when the carbon-to-nitrogen (C/N) was set at 10:1 or 15:1, the optical density at OD₆₀₀ values for the strain were 1.25 ± 0.01 and 1.13 ± 0.01 (P < 0.05), respectively, with NH₄⁺-N utilization efficiency reaching 100%, with a utilization rate of 4.15 mg/L/h. At a C/N ratio of 5, the OD₆₀₀ was 1.14 ± 0.01, and the NH₄⁺-N utilization efficiency was 95.63%. This utilization efficiency was higher than the 90% NH₄⁺-N utilization efficiency reported for Pseudomonas sp. M-33 [53] at a C/N ratio of 5, as well as the average NH₄⁺-N utilization rates of 3.70 mg/L/h observed for Bacillus thuringiensis strain WXN-23 at the same C/N ratio [73]. As the C/N ratio increased, the accumulation of NO₃^--N at the end of the culture period rose from 0.04 ± 1.54 mg/L at a C/N ratio of 5 to 16.16 ± 0.57 mg/L, indicating enhanced nitrification capacity. When the C/N ratio reached 20, the NH₄⁺-N utilization efficiency decreased to 98.70%. This reduction could be attributed to excess organic matter, which may directly interact with the enzyme structure, affecting enzyme activity and inhibiting both strain growth and nitrification activity [67]. Based on the NH₄⁺-N utilization efficiency, growth, and nitrification efficiency, the optimal C/N ratio range for subsequent experiments was determined to be between 5 and 15.

As shown in Fig. 8(c), At pH levels of 7 and 8, the optical density at OD₆₀₀ values were 1.24 and 1.01, respectively, significantly higher than those observed at pH levels 5 and 6 (OD₆₀₀ = 0.36 ± 0.08 and 0.64 ± 0.01, P < 0.05). At pH 7, the utilization efficiency of NH₄⁺-N reached 100%, and within the pH range of 7–9, the NH₄⁺-N utilization efficiency varied from 90.85 to 100%. This performance surpassed that of Aeromonas sp. HN-02 [74], which exhibited a utilization efficiency of NH₄⁺-N 82.93–90.20% at pH levels of 7–9. When the pH was 5 and 10, the accumulated of NO₃⁻-N concentrations were 53.74 ± 0.96 mg/L and 12.35 ± 0.54 mg/L, respectively. These results suggest that strain GW-E demonstrates strong nitrification performance and a broad pH tolerance range. However, at pH 6, the NH₄⁺-N utilization efficiency was 69%, significantly lower than at other pH values. Considering bacterial growth (OD₆₀₀), NH₄⁺-N utilization efficiency, and the stable accumulation of NO₃⁻-N and NO₂⁻-N at pH 7–9, the results indicate that the strain demonstrates superior nitrification performance across a wide pH spectrum.

As shown in Fig. 8(d). At 55 °C, the OD₆₀₀ of strain GW-E was 1.24 ± 0.06, indicating the highest NH₄⁺-N utilization efficiency of 100%. In contrast, at the same temperature, Anoxybacillus contaminans HA [75] demonstrated an NH₄⁺-N utilization efficiency of only 71%. When the cultivation temperature was increased to 60 °C, the NH₄⁺-N utilization efficiency reached 92.85%, with cumulative production of NO₃⁻-N and NO₂⁻-N of 16.54 ± 1.64 mg/L mg/L and 15.41 ± 0.40 mg/L mg/L, respectively. These values were significantly higher than those observed for the thermotolerant strains Brevibacillus agri N2 (NH₄⁺-N utilization efficiency 43.6%) and Gordonia paraffinivorans N52 (NO₃⁻-N accumulation 4.2 ± 0.29 mg/L at 60 °C). The greater accumulation of NO₃⁻-N and NO₂⁻-N, along with the enhanced NH₄⁺-N utilization efficiency in the strain GW-E, compared to previously reported thermophilic heterotrophic nitrifying strains, suggested that strain GW-E exhibits strong adaptability to temperature variations [76]. In summary, the strain demonstrated a greater capacity for ammonia nitrogen utilization within a temperature range of 50–60 °C, a pH level between 7 and 9, and a carbon-to-nitrogen (C/N) ratio of 5 to 15. Subsequently, response surface methodology will be employed to further optimize the cultivation conditions for strain GW-E.

Optimization of cultivation conditions using response surface methodology

Owing to the complexity of environmental conditions, microorganisms have varying abilities to grow and adapt to different environmental factors, necessitating an optimal combination of these elements. The most direct indicator of heterotrophic nitrification is NH₄⁺-N utilization efficiency. Because nitrate and nitrite are unstable and can easily be converted into one another, a two-dimensional model was established to describe the interaction between independent variables and NH₄⁺-N utilization efficiency using Design-Expert software, and a quadratic polynomial regression analysis was conducted on the data presented in Table 4. A multiple regression equation was derived for temperature (A), pH (B), and C/N ratio (C) in relation to the NH₄⁺-N utilization efficiency (Y). The equation is as follows:

Table 4.

Analysis of variance (ANOVA) of quadratic parameters for NH₄⁺-N utilization efficiency

Source	Sum of Squares	df	Mean Square	F-Value	P-value	significant
Model	2374.90	9	263.88	29.44	< 0.0001	**
A-temperature	119.58	1	119.58	13.34	0.0082	**
B-pH	175.31	1	175.31	19.56	0.0031	**
C-C/N	736.51	1	736.51	82.16	< 0.0001	**
AB	4.84	1	4.84	0.5399	0.4863	−−
AC	178.89	1	178.89	19.96	0.0029	**
BC	1.60	1	1.60	0.1785	0.6853	−−
A2	760.68	1	760.68	84.86	< 0.0001	**
B2	15.94	1	15.94	1.78	0.2241	−−
C2	301.60	1	301.60	33.65	0.0007	**
Residual	62.75	7	8.96	−−	−−	−−
Lack of Fit	21.27	3	7.09	0.6836	0.6070	−−
Pure Error	41.48	4	10.37	−−	−−	−−
Cor Total	2437.65	16	−−	−−	−−	−−

* indicates p < 0.05; * * indicates p < 0.01

Y(%) = 96.012 + 3.86625A + 4.68125B + 9.595C + 1.1AB – 6.6875AC – 0.6325BC – 13.441A² – 1.946B² – 8.4635C².

The correlation coefficient (R²) was 0.9743. ANOVA for the model indicated that the three primary factors influenced NH₄⁺-N utilization efficiency in the following order of significance : C/N ratio > pH > temperature. As shown in Fig. 9, the C/N ratio had a highly significant effect on NH₄⁺-N utilization efficiency, whereas the effects of temperature and pH were comparatively smaller. The final optimized conditions predicted by the response surface model were a temperature of 54.9℃, a pH of 8, and a C/N ratio of 12.20, resulting in NH₄⁺-N utilization efficiency of 100%.

Fig. 9.

The response surface plots and corresponding contour plots of NH₄⁺-N utilization efficiency of strain GW-E. NH₄⁺-N utilization efficiency as a function of Temperature and pH (a, b); NH₄⁺-N utilization efficiency as a function of pH and C/N ratio (c, d); NH₄⁺-N utilization efficiency as a function of C/N ratio and temperature (e, f)

Response surface validation experiment

The regression equation model was analyzed, and following response surface optimization, the optimal cultivation conditions were established as follows: a carbon-to-nitrogen (C/N) ratio of 15:1, a temperature of 54 °C, and an initial pH of 8. Under these conditions, the NH₄⁺-N utilization efficiency of the strain reached 99.928%, while the assimilation rate after optimization was 56.19%, representing an increase of 16.15% compared to the rate before optimization. After conducting three repeated trials, the relative error between the measured value and the model’s predicted value was 0.008%, indicating that the model’s simulation performance was excellent. The optimal temperature of 54 °C and the slightly alkaline pH of the strain align with the high-temperature phase of composting (45–60 °C), and the strain maintained an NH₄⁺-N utilization rate exceeding 98.7% under a high C/N ratio (15–20) [11, 77].

Advantages and application prospects of strain GW-E

The thermotolerant strain GW-E exhibits exceptional nitrogen utilization efficiency under high-temperature conditions (55 °C), achieving complete ammonium removal (100% NH₄⁺-N utilization efficiency) and significant nitrate/nitrite reduction (21.44% NO₃⁻-N and 51.68% NO₂⁻-N utilization efficiency, respectively) following optimization. These performance metrics substantially exceed those of conventional heterotrophic nitrifiers. Strains such as Acinetobacter indicus CZH-5, Acinetobacter oleivorans AHP123, and Sneathiella aquimaris 216LB-ZA1 exhibit lower utilization rates within the cultivation period compared to this strain [40, 78, 79]. Particularly, research on thermophilic heterotrophic nitrifying strains is scarce, as shown in Table 5. The mechanisms underlying high-temperature tolerance remain limited to phenotypic validation. Therefore, this study infers the nitrogen utilization pathways of strain GW-E through multidimensional approaches. The strain utilizes three distinct nitrogen transformation pathways—assimilatory nitrate reduction, nitrification, and ammonia assimilation—with approximately 40.04% of nitrogen being incorporated into cellular biomass, thereby effectively minimizing nitrogen loss. Genomic characterization indicates that its thermotolerance mechanism involves the coordinated regulation of the gyrB, rbfA, and punA genes, along with a heat shock protein protection system, which collectively maintain cellular integrity and nitrogen metabolic functions under thermal stress [69]. The optimal growth conditions (55 °C, C/N ratio of 20–35) align perfectly with thermophilic composting parameters, addressing the critical limitation of mesophilic bacterial inactivation during high-temperature composting phases.

Table 5.

Comparison of nitrogen utilization capacity and mechanism of high-temperature heterotrophic nitrifying bacteria

Bacterial genus name	Separate sources	NH4+-N utilization efficiency	NO3−-N utilization efficiency	NO2−-N utilization efficiency	Enzyme activity assay	Genome wide analysis	Assimilation efficiency	optimal temperature	References
Brevibacillus Agri N2	Sewage sludge composting	45.47% (99.64 mg/L)	-	76.77%(55.20 mg/L)	AMO 15.19 ± 0.91 HAO 12.29 ± 0.41 NXR 21.86 ± 1.40(U/L)	-	-	60℃	[16]
Gordonia paraffinivorans N52	Sludge composting	51.8%	-	-	AMO 18.4 ± 0.7 HAO 15.9 ± 0.5 NXR 28.9 ± 1.5 (U/L)	-	53.0%	60℃	[17]
Anoxybacillus contiminans HA	-	71.0%	74.7%	-	-	-	-	55℃	[80]
Chelatococcus daeguensis TAD1	-	67.7%	-	-	-	-	-	50 °C	[81]
Aeribacillus pallidus sp. GW-E	High-temperature composting	100%	21.44%	51.68%	AMO 0.32 ± 0.11 HAO 0.04 ± 0.01 NAR 0.05 ± 0.00 NIR 0.04 ± 0.00 GS 1.014 ± 0.030 GDH 1.114 ± 0.090 GOGAT1.611 ± 0.061 (U/mL)	Assimilation of nitrate reduction pathways, nitrification pathways, and ammonia assimilation pathways	40.04%	55–60℃	This study

The strain GW-E demonstrates significant potential for aerobic composting applications due to its exceptional thermotolerance, which enables efficient nitrification during the thermophilic phase (45–60℃). Comparative studies highlight its superior performance compared to established thermophilic nitrifiers: Gordonia paraffinivorans N52 achieves 53.0% nitrogen assimilation at 50 °C, enhancing NO₃⁻-N accumulation while reducing NH₃ emissions and total nitrogen loss [17]. In contrast, Bacillus stearothermophilus, which has an optimal growth temperature of 55–65 °C, increases nitrifier populations and decreases NH₃ release [82]. The optimal growth temperature of GW-E (55 °C) aligns perfectly with the thermophilic composting range (45–60 °C) [83], suggesting its potential for enhanced nitrification performance while concurrently reducing NH₃ and N₂O emissions during high-temperature phases. This capability supports both environmental protection and the valorization of livestock waste. Furthermore, GW-E’s compatibility with optimal composting carbon-to-nitrogen (C/N) ratios (20–35) [84], which critically influence microbial activity and organic matter decomposition, confirms its practical applicability in composting systems. Future research could further elucidate the molecular synergistic mechanisms between heat shock proteins and nitrogen metabolism enzymes, providing theoretical support for the large-scale application of this bacterium in aerobic composting of livestock and poultry. This would contribute to the dual benefits of environmental protection and resource recycling.

Conclusions

This study successfully isolated and characterized the thermotolerant heterotrophic nitrifying bacterium Aeribacillus pallidus sp. GW-E, which demonstrated remarkable nitrogen transformation capabilities at 55 °C. The strain exhibited utilization efficiencies of 87.42%, 21.44%, and 51.68% for NH₄⁺-N, NO₃⁻-N, and NO₂⁻-N, respectively, with NH₄⁺-N utilization efficiency reaching 100% after response surface optimization. Enzymatic assays revealed significantly higher specific activities of glutamine synthetase (1.014 ± 0.030 U/mg), glutamate dehydrogenase (1.114 ± 0.090 U/mg), and glutamate synthase (1.611 ± 0.061 U/mg) compared to other nitrogen metabolic enzymes. The strain utilized multiple nitrogen transformation pathways, including ammonium assimilation (40.04% nitrogen incorporation into biomass), assimilatory nitrate reduction (key gene nirA), and nitrification (NH₄⁺→NO₂⁻/NO₃⁻). Genomic analysis identified heat shock genes (hslO, groES) that contribute to its thermotolerance. The optimal growth conditions (C/N ratio = 15, 54 °C, pH = 8) closely matched typical thermophilic composting parameters. These findings provide a high-quality microbial resource for nitrogen retention during thermophilic composting and establish a theoretical foundation for developing efficient microbial inoculants in organic waste recycling applications.

Authors’ contributions

“Z and M,Q,G wrote the main manuscript text and Gand X prepared all figures.S,W,L modify manuscript Conceptualization, ValidationAll authors reviewed the manuscript.”

Funding

This work was financially supported by National Natural Science Foundation of China (32160756); Gansu Science and Technology Innovation Guidance Plan - Eastern-Western Regional Sci-Tech Cooperation Initiative (25CXNA029); Fuxi Foundation of Gansu Agricultural University (GAUfx-04J03); National Natural Science Foundation of Gansu Province (24JRRA644); the Discipline Team Project of Gansu Agricultural University (GAU-XKMB2022-20); Research and Application of Supporting Technologies for Gansu Pig Industry (GNKJ-2024-46).

Data availability

The datasets generated and/or analysed during the current study are available from the following link: https://www.ncbi.nlm.nih.gov/search/all/?term=PRJNA1224725.

Declarations

Ethics approval and consent to participate

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Not applicable.

Competing interests

The authors declare no competing interests.