Advances in Heterotrophic Nitrification–Aerobic Denitrification: Physiological Characteristics, Environmental Responses, and Nitrogen Metabolic Pathways

Abstract

Heterotrophic nitrification–aerobic denitrification (HNAD) bacteria enable nitrogen removal under aerobic conditions and have attracted increasing attention in wastewater treatment. Although numerous studies have enriched HNAD strains and explored their nitrogen removal performance, the environmental responses and metabolic mechanisms underlying nitrogen transformation remain insufficiently evaluated. In this review, the ecological distribution and physiological characteristics of HNAD bacteria are summarized, followed by a systematic elaboration of the effects of nitrogen sources, carbon sources, C/N ratio, and environmental factors (e.g., temperature, pH, dissolved oxygen and salinity) on nitrogen removal and transformation performance. In addition, nitrogen depletion from the aqueous phase is likely to reflect both conversion to gaseous nitrogen products and assimilatory uptake into biomass, which may account for a non-negligible proportion of the apparent nitrogen removal in some strains. Accordingly, the nitrogen transformation mechanisms, encompassing nitrite/nitrate-mediated pathways, proposed hydroxylamine-associated pathways, and nitrogen assimilation, are critically examined, with attention to the strength and limitations of the available evidence. Furthermore, the regulatory roles of intracellular redox balance and electron allocation in mediating the interplay among nitrification, denitrification, and cellular respiration are addressed. Finally, the key challenges and future prospects of HNAD research and applications are highlighted. This review provides mechanistic insights into the metabolic versatility of HNAD bacteria and supports the development and evaluation of nitrogen removal strategies in advanced biological nitrogen removal systems.

1. Introduction

In recent years, water pollution caused by excessive nitrogen emissions, including municipal sewage, industrial wastewater, livestock breeding wastewater, and agricultural non-point source pollution, has emerged as a critical environmental concern and attracted increasing attention worldwide [1,2]. Among inorganic nitrogen pollutants, ammonia nitrogen warrants particular attention due to its greater acute toxicity to aquatic organisms compared with nitrate. Its toxicity mainly depends on the equilibrium between ammonium (NH₄⁺) and the toxic free ammonia (NH₃), which varies with pH and temperature [3]. Effective ammonium removal is therefore particularly important for mitigating the immediate ecological risks of nitrogen-rich effluents. Biological nitrogen removal (BNR) has been widely applied due to the advantages of high nitrogen removal efficiency, low cost and low secondary pollution when compared to the physical-chemical nitrogen removal processes [4,5]. In the mainstream wastewater treatment plants (WWTPs), BNR processes mainly include aerobic nitrification and anoxic denitrification, where ammonia is oxidized to nitrate by autotrophic nitrifiers under aerobic conditions and nitrate is subsequently reduced to nitrogen gas by heterotrophic microorganisms under anoxic conditions [6].

Nowadays, several new nitrogen biotransformation processes have emerged to overcome the deficiencies in the conventional BNR systems, mainly including simultaneous nitrification and denitrification (SND), endogenous denitrification (ED), anaerobic ammonium oxidation (Anammox), partial nitrification/denitrification (PN/D), sulfur autotrophic denitrification (SADN), hydrogen autotrophic denitrification (HD) and Heterotrophic nitrification–aerobic denitrification (HNAD) [7]. Among these processes, HNAD enables nitrification and denitrification to occur within a single organism/system under aerobic conditions [7,8], which provides opportunities to integrate nitrogen and organic matter removal in a single aerated system and therefore eliminates the need for separate oxic and anoxic treatment stages [8]. Heterotrophic nitrification (HN) generally refers to the oxidation of reduced nitrogen compounds, including ammonium and certain organic nitrogen compounds, to nitrite and/or nitrate by heterotrophic microorganisms [9], and this process may not be directly coupled to energy conservation for driving ammonia oxidation. Aerobic denitrification (AD), by contrast, is the reduction of nitrate or nitrite to gaseous nitrogen products under aerobic conditions [10], where organic carbon may serve as an electron donor, an energy source, and a carbon source for biomass synthesis [11]. At the enzymatic level, ammonia monooxygenase (AMO) and hydroxylamine oxidase (HAO) activities have been reported in individual HNAD strains [12], while enzyme purification and characterization in Thiosphaera pantotropha identified a hydroxylamine oxidase enzyme distinct from those of autotrophic nitrifiers [13]. Aerobic denitrification mainly involves nitrate reductases (Nar/Nap), nitrite reductases (NirK/NirS), nitric oxide reductase (Nor), and nitrous oxide reductase (NosZ), and the activity is linked to carbon availability and cellular redox balance through electron partitioning between oxygen respiration and nitrogen oxide reduction [14,15]. However, the involvement of canonical nitrification enzymes or genes cannot be assumed for all HNAD strains. For example, genomic analysis of Pseudomonas stutzeri UFV5 identified denitrification genes but failed to identify genes encoding the canonical autotrophic nitrification enzymes [16], which suggests the potential existence of alternative ammonia-oxidizing functional genes or pathways in HNAD strains.

The occurrence of HNAD has overcome the limitations of the conventional BNR process. Compared with autotrophic nitrifiers, HNAD bacteria have been reported to grow faster, exhibit greater adaptability to environmental fluctuations, and utilize organic substrates more efficiently [17,18]. By combining heterotrophic nitrification and aerobic denitrification, HNAD enables simultaneous nitrogen and COD removal in a single aerobic system [19], although the performance strongly depends on bacterial strain, nitrogen substrate, carbon source and availability, and environmental conditions, particularly dissolved oxygen (DO) [16,19]. Compared with anaerobic ammonium removal (e.g., anammox) systems, HNAD offers distinct process advantages, particularly for wastewater containing organic carbon [20]. For example, HNAD enables simultaneous nitrification and denitrification, providing simpler operation, faster start-up, and greater adaptability to environmental fluctuations, as well as enhanced robustness toward organic carbon loading [21,22]. These characteristics offer potential advantages for treating wastewaters containing both ammonium and biodegradable organic matter and may reduce reactor footprint and operational complexity.

Currently, various HNAD bacteria have been reported in multiple environments, including landfill leachate [23,24], wastewater treatment plants [25,26], activated sludge [17], soil [27], rivers [28], marine aquaculture systems [29], and coking wastewater [30], with reported isolates belonging to genera such as Acinetobacter, Bacillus, Cuprobacter, Halomonas, Klebsiella, Marinobacter, Pseudomonas, and Photobacterium [31]. However, these HNAD strains exhibit distinct responses to environmental conditions and often require specific physiological settings for optimal performance. In addition, the colonization potential and competitive fitness of individual strains within complex microbial communities remain poorly understood. Recent studies have suggested that different HNAD bacteria may adopt distinct nitrogen metabolism strategies, reflecting the complexity of HNAD pathways. Although several representative metabolic pathways of HNAD have been proposed, the exact nitrogen transformation routes and their regulatory mechanisms remain to be fully elucidated. Accordingly, a systematic understanding of bacteria-mediated HNAD processes is still needed, particularly regarding the nitrogen transformation pathways and the underlying metabolic mechanisms.

Previous reviews have summarized HAND strain diversity, environmental influences, metabolic pathways, and key enzymes [32], as well as the coupling of organic carbon metabolism and nitrogen conversion through electron transport [8]. Building on these syntheses, integrating strain-specific physiological responses with experimental evidence for nitrogen transformation pathways is essential to explain variation in reported HNAD performance. Recent findings have also highlighted the need to reassess proposed nitrogen removal pathways: isotope-based and transcriptomic analyses of Alcaligenes faecalis identified hydroxylamine as the primary nitrification product and detected no denitrification under the conditions tested [10]. An updated synthesis is therefore needed to link environmental responses to strain-specific metabolic evidence and distinguish shared physiological features from mechanisms established only in individual isolates.

This review focuses on HNAD bacterial isolates characterized in pure culture and comprehensively summarizes their distribution and physiological characteristics. The key environmental factors affecting the performance of HNAD bacteria are also systematically evaluated. Furthermore, the HNAD metabolic mechanisms are examined in terms of electron transfer and nitrogen transformation. Building on previous reviews, particular attention is given to the relationships among strain characteristics, cultivation conditions, and proposed nitrogen transformation pathways. Finally, critical challenges and future prospects for HNAD research and application are highlighted. This review aims to provide a clear framework for understanding the physiological characteristics and metabolic mechanisms of HNAD strains, which could thereby support the development and optimization of efficient biological nitrogen removal processes in wastewater treatment systems.

2. Distribution and Enrichment of HNAD Bacteria

2.1. The Discovery and Progress

In 1983, Robertson and Kuenen reported the isolation of Thiosphaera pantotropha, later reclassified as Paracoccus pantotrophus, from a desulfurizing and denitrifying effluent-treatment system [33]. A subsequent study demonstrated that this bacterium could simultaneously perform heterotrophic nitrification and aerobic denitrification in aerobic chemostat cultures [34]. Since then, extensive research has focused on the isolation of various HNAD strains and evaluation of their nitrogen removal performance under different environmental conditions [31,35]. Table 1 summarizes 22 taxonomic genera of representative HNAD strains isolated in pure culture, together with their isolation sources and characterization methods. These strains have been experimentally shown to exhibit both HN and AD activities to remove ammonium, nitrite, and nitrate effectively.

Table 1. Representative HNAD-capable bacterial isolates, their isolation sources, and the methods used to assess their nitrogen-transforming activities.

Genus	Representative Strain	Isolation Source	Nitrogen and Carbon Source	Enrichment Conditions	Methods for Assessing HNAD Activity	Reference
Pseudomonas	Pseudomonas putida strain LX1	Activated sludge	NH4+-N; sodium succinate hexahydrate	30 °C; 160 rpm	Aerobic NH4+/NO3−-N removal assays; AMO and HAO activity assays	[36]
Acinetobacter	Acinetobacter oleivorans AHP123	Activated sludge	NH4+-N; sodium succinate	30 °C; 200 rpm; pH 7.3; C/N 20	NH4+/NO3− removal assays; nitrogen balance; functional gene annotation and RT-qPCR	[37]
Bacillus	Bacillus sp. L2	Aeration tank water samples	NH4+-N; sodium acetate	30 °C; 160 rpm; pH 7.0	NH4+/NO3−/NO2− removal assays; PCR amplification of HNAD-related genes	[38]
Klebsiella	Klebsiella sp. TN-10	Tannery wastewater	NH4+-N; sucrose	30 °C; 150 rpm; pH 7.0	NH4+/NO3−/NO2− removal assays; TN monitoring; AMO, nitrate reductase and Nir activity assays	[39]
Halomonas	Halomonas venusta SND-01	Marine sediment	NH4+-N; sodium citrate	30 °C; 120 rpm; pH 7.5–8.5; NaCl (30 g/L)	NH4+/NO3−/NH2OH/NO2− removal assays; genome annotation; AMO, HAO, Nir and Nar activity assays	[40]
Alcaligenes	Alcaligenes ammonioxydans sp. HO-1	Wastewater treatment sludge	Not reported	Not reported	NH4+/NO2− conversion assays; gaseous product analysis; 15N tracing; denitrification inhibition assay; functional cloning of dnf gene	[41]
Vibrio	Vibrio sp. LY1024	Not report	NH4+-N and NO2−-N; sodium citrate	pH 7.2	NH4+/NO3−/NO2− removal assays; nitrogen intermediate monitoring	[42]
Zobellella	Zobellella endophytica W14	Soil of Ulansuhai Wetland	NO3−-N; sodium succinate	30 °C, 150 rpm; pH 7.0–7.2; NaCl (50 g/L)	Nitrogen removal assays; nitrogen balance; PCR amplification of hao, napA, nirS and nosZ	[43]
Gordonia	Gordonia amicalis UFV4.	Activated sludge	Not reported	Not reported	NH4+ removal assays; gaseous product analysis; nitrogen balance	[44]
Cupriavidus	Cupriavidus sp. S1	Coking wastewater	NH4+-N; sodium pyruvate	pH 7.0; 30 °C; 120 rpm	NH4+/NO3−/NO2− removal assays; TN monitoring; N2 detection	[30]
Thiosphaera	Thiosphaera pantotropha LMD 82.5	Sulphide-oxidizing–denitrifying treatment system	NH4+-N and NO3−-N; acetate and thiosulfate	pH 8.0; 37 °C; dilution rate 0.03 h−1	Nitrogen balance; O2 uptake assays; AMO and HAO activity assays	[45]
Aeromonas	Aeromonas sp. HN-02	Activated sludge	Not reported	Not reported	NH4+/NO3−/NO2− removal assays; TN monitoring	[46]
Providencia	Providencia rettgeri YL	MBR reactor	Peptone and beef extract	NaCl; 30 °C; 120 rpm; 24 h	15N tracing; N2/N2O isotope analysis; nitrogen balance	[47]
Serratia	Serratia sp. LJ-1	Membrane bioreactor	NH4+-N; glucose and peptone	pH 7.0–8.0; 30 ± 2 °C 100 rpm	NH4+ removal and NO3−/NO2− production assays; nitrogen-species monitoring	[48]
Streptomyces	Streptomyces mediolani EM-B2	Not reported	Not reported	Not reported	Nitrogen removal assays; nitrogen balance; inhibition assays; AMO, HAO, NR and Nir activity assays	[49]
Diaphorobacter	Diaphorobacter sp. PD-7	Activated sludge	NH4+-N; phenol	pH 7.2–7.4; 30 °C; 180 rpm	NH4+/NO3− removal assays; nitrogen intermediate monitoring; HAO, Nar and Nir activity assays	[50]
Delftia	Delftia tsuruhatensis SDU2	Waste treatment pond of pig farm	Acetamide	30 °C	NH4+/NO3−/NO2− removal assays; TN and biomass-N measurements; PCR amplification and RT-qPCR of HNAD-related functional genes	[51]
Achromobacter	Achromobacter sp. GAD-3	Sediment and sewage from the Longfeng Wetland	NH4+-N and NO3−-N; sodium acetate	pH 7.0; 30 °C; 150 rpm	Nitrogen-species monitoring; gaseous product analysis; nitrogen balance; Nar, Nir, Nor and N2or activity assays; RT-qPCR	[52]
Janthinobacterium	Janthinobacterium sp. J1-1	Wastewater	NH4+-N; sodium succinate	pH 7.0 ± 0.2; 8 °C; 100 rpm	NH4+/NO3−/NO2− removal assays; TN monitoring; genome annotation; PCR amplification of HNAD-related functional genes	[53]
Ochrobactrum	Ochrobactrum anthropic LJ81	Domestic sludge	NO3−-N; sucrose	30 °C; 150 rpm	NH4+/NO3−/NO2− removal assays; nitrogen balance; chlorate inhibition assays; PCR amplification of denitrification gene	[54]
Chryseobacterium	Chryseobacterium sp. R31	Abattoir wastewater	NH4+-N; glucose	pH 6.8–7.5, 120 rpm	NH4+/NO3−/NO2− removal assays; nitrogen balance estimation; chlorate inhibition; denitrification gene PCR	[55,56]
Rhodococcus	Rhodococcus sp. CPZ24	Swine wastewater ponds	NH4+-N; sodium succinate hexahydrate	30 °C; pH 7.2; 120 rpm	NH4+/NO3− removal assays; intracellular-N measurement; N2/N2O detection	[57]

HNAD strains have been isolated from a wide range of environments, including activated sludge, wastewater, soil, and freshwater lakes [32], as shown in Figure 1. Organic carbon utilization and aerobic nitrogen transformation provide a physiological basis for HNAD bacteria to exploit habitats containing suitable carbon and nitrogen substrates [58]. In some isolates, tolerance to a wide range of salinity and pH may further facilitate their occurrence or persistence in particular habitats [40]. However, enrichment and screening inherently select for strains capable of both growth and nitrogen removal under the selected substrate and cultivation conditions [59]. Therefore, the isolates obtained may not fully represent the natural HNAD community.

Figure 1. Distribution of isolation environments of HNAD bacteria.

Moreover, numerous studies have reported HNAD-associated genera in specific engineered treatment systems. For example, in marine aquaculture wastewater treatment systems, Bacillus and Halomonas were identified as keystone genera associated with HNAD activity [60]. In wastewater treatment plants, the putative HNAD-associated genera Denitratisoma, Dechloromonas, and Comamonas showed their highest combined relative abundance in the A/O process, accounting for 7.69% of the microbial community [61]. In addition, Acinetobacter was reported to dominate the putative HNAD-associated community in MBR sewage treatment systems [62]. These community profiles suggest the occurrence of potentially relevant taxa. However, the contribution of these taxa to HNAD still requires functional verification. The reported isolation sources in Table 1 provide a basis for selecting samples for further enrichment, purification, and characterization.

2.2. The Enrichment and Purification

In existing studies, HNAD bacteria are generally isolated through selective enrichment and subsequent purification [63,64]. Most previously reported HNAD strains have been enriched under aerobic conditions in chemically defined media, where ammonium, nitrite, or nitrate serves as the sole/mixture nitrogen source, with an organic carbon source provided to support heterotrophic metabolism [65,66,67]. These conditions impose selective pressure by favoring microorganisms capable of nitrogen transformation under simultaneous aerobic, heterotrophic conditions. Nevertheless, evidence of nitrogen removal obtained solely during enrichment is insufficient to establish an HNAD phenotype, as the disappearance of inorganic nitrogen may also reflect assimilation or other nitrogen-transforming processes [68]. In addition, HNAD activity may be attributed to a single strain or to community-level cooperation [69,70]. Community-level nitrogen removal alone is, therefore, insufficient to establish that individual members perform both heterotrophic nitrification and aerobic denitrification, nor does it indicate which members contribute to these processes. After purification, strain-specific assessment should distinguish ammonium assimilation from heterotrophic nitrification and evaluate nitrate and/or nitrite reduction under aerobic conditions [27]. Analysis of nitrogen transformation products and nitrogen mass balances [71,72], supplemented by enzyme assays or genetic evidence, can further help substantiate the proposed pathways.

A wide range of HNAD strains affiliated with genera such as Alcaligenes, Pseudomonas, and Acinetobacter have been successfully isolated and purified as shown in Table 1. For example, Li et al. enriched HNAD bacteria from activated sludge in an MBR wastewater treatment system and isolated 53 HNAD strains, among which Acinetobacter was the dominant genus [62]. Previous studies have demonstrated that enrichment under specific selection pressures (e.g., high salinity, high nitrogen loading and low C/N ratio) may favor the isolation of HNAD strains capable of growth and nitrogen removal under the selected conditions. For instance, enrichment under saline conditions yielded halotolerant strains such as Halomonas venusta TJPU05, which achieved over 80% removal of multiple nitrogen species (NH₄⁺, NO₃⁻ and NO₂⁻) at high salinity (8%) [73]. Alcaligenes faecalis TF-1, isolated from landfill leachate with a strategy of gradual increase in nitrogen loading, was reported to tolerate an extremely high concentration of NH₄⁺-N at 2000 mg N L⁻¹ and achieved 94.6% NH₄⁺-N removal under optimized conditions [74]. Similarly, Acinetobacter sp. WZ-1, obtained under low C/N enrichment conditions, removed 47–88% of NH₄⁺-N at C/N ratios of 4–8 [75]. Collectively, these examples suggest that tailoring enrichment conditions to the target environment may help obtain HNAD strains with the desired physiological and nitrogen-removal characteristics. The microbial responses of HNAD strains to specific environmental factors are further discussed in Section 4.

3. Physiological Characteristics of HNAD Bacteria

3.1. Morphological Diversity of Reported Isolates

Bacteria capable of HNAD belong to diverse taxonomic groups, and this taxonomic diversity is usually reflected in varying morphological features. Among the reported isolates, rod-shaped and short-rod forms predominate, although coccoid and pleomorphic forms have also been documented, which is dependent on taxonomic affiliation and cultivation conditions [64]. In addition, reported cells typically measure 0.5–1.0 μm in width and 1.0–3.0 μm in length [27].

3.2. Kinetic Parameters

The maximum specific growth rate (${\mu }_{\max }$) and apparent half-saturation coefficient ($K_s$) are commonly used to describe the growth kinetics of HNAD bacteria. In a Monod growth model, ${\mu }_{\max }$ represents the maximum specific growth rate under the specified conditions, whereas $K_s$ is the concentration of the growth-limiting substrate at which the growth rate reaches half of ${\mu }_{\max }$ [76]. $K_s$ is an apparent, model-dependent coefficient specific to the substrate, process, and experimental conditions, rather than a direct measure of intrinsic biochemical affinity [77]. Apparent $K_s$ values estimated for ammonium, nitrite, nitrate, or other growth-limiting substrates are therefore not directly interchangeable. Reported ${\mu }_{\max }$ values for the HNAD strains summarized in Table 2 range from approximately 0.18 to 0.60 h⁻¹, with variation reflecting both strain characteristics and cultivation conditions. Comparisons of kinetic parameters require sufficiently comparable substrates, processes, and experimental conditions. These growth parameters help characterize growth potential for the specific HNAD strains under their respective cultivation conditions, but may not establish a competitive advantage in the mixed communities.

Table 2. Comparison of ${\mu }_{\max }$ between HNAD bacteria and autotrophic nitrifiers.

Microorganism Type	Representative Strains	Nitrogen Substrate	${\mathit{\mu }}_{\mathbf{max}}$ (h−1)	Reference
HNAD bacteria	Alcaligenes faecalis No. 4	NH4+-N	0.20	[64]
	Agrobacterium sp. LAD9	NH4+-N	0.32	[78]
	Achromobacter sp. GAD3/GAD4		0.31/0.38
	Bacillus spp.	NH4+-N	0.43–0.55	[65]
	Paracoccus denitrificans ISTOD1	NH4+-N	0.52	[79]
	Cupriavidus sp. S1	NH4+-N	0.186	[30]
	Klebsiella pneumoniae CF-S9	NH4+-N	0.55	[80]
	Ochrobactrum anthropic LJ81	NH4+-N, NO3−-N, NO2−-N	0.36, 0.324, 0.315	[54]
	Pseudomonas stutzeri KTB	NH4+-N, NO3−-N, NO2−-N	0.56, 0.42, 0.45	[81]
	Thiosphaera pantotropha	NH4+-N, NH4+-N+NO3−-N, NO3−-N	0.28–0.45, 0.34–0.58, 0.25–0.49	[34]
Autotrophic nitrifiers (AOB)	Nitrosomonas europaea	NH4+-N	0.03–0.05	[34]
	Nitrosomonas mobilis Ms1	NH4+-N	0.05–0.07	[82]

${\mu }_{\max }$: maximum specific growth rate; Values were reported in the literature and may vary with temperature, substrate type, and cultivation conditions.

3.3. Nitrogen Source Selectivity

HNAD bacteria can utilize diverse inorganic nitrogen sources, including NH₄⁺-N, NO₂⁻-N and NO₃⁻-N under aerobic conditions [32]. However, substrate preferences vary among strains and can influence growth and nitrogen removal performance (Table 3) [55]. Generally, NH₄⁺ is more readily utilized than NO₂⁻ or NO₃⁻ [83], although substrate preference may vary depending on microbial species and environmental conditions.

Table 3. Nitrogen source preference and removal characteristics of representative HNAD bacteria.

Strain	Nitrogen Preference	Nitrogen Removal Rates (mgN·L−1·h−1) or Efficiency (%)	Nitrogen Transformation Traits	Reference
Acinetobacter sp. ND7	NH4+ > NO2− > NO3−	NH4+ 5.7; NO2− 4.2; NO3−: 3.6	NO2− accumulation	[70]
Lysinibacillus fusiformis B301	NH4+ > NO3− > NO2−	NH4+ 2.1; NO3− 1.6; NO2− 1.4	83.25% NH4+-N to N2	[84]
Halomonas sp. 5505	NH4+ prior to NO3−/NO2−	NH4+ 100%; NO3− 94.1%; NO2− 74.4%	Assimilation-dominated	[85]
Hydrogenophaga sp. H7	NH4+ > NO3−	NH4+ 1.4; NO3− 0.9;	NH4+-N removal inhibited by NO3−/NO2− addition	[86]
	NH4+ > NO2−	NH4+ 1.6; NO2− 2.3
Paracoccus denitrificans HY-1	NH4+ prior to NO3−/NO2−	NH4+ 10.7; NO3− 22.1; NO2− 27.8	NH4+-N enhanced the removal of NO3−-N and NO2−-N	[87]
Pseudomonas putida ZN1	NH4+ prior to NO3−/NO2−	NH4+ 94.5%; NO3− 86.1%; NO2− 71.6%	Preferential NH4+-N oxidation followed by denitrification without intermediate accumulation	[88]

When NH₄⁺-N was provided as the sole nitrogen source, most HNAD strains often exhibited comparable or improved nitrogen removal performance and growth behavior when compared with the oxidized nitrogen sources (e.g., NO₂⁻-N and NO₃⁻-N), although this response was highly strain-dependent. This preference may partly reflect that NH₄⁺ can be directly utilized in HN or directly assimilated into biomass, whereas oxidized nitrogen forms require additional reduction steps before incorporation into cellular biomass [89]. For example, Pseudomonas stutzeri YG-24 exhibited average NH₄⁺-N, NO₂⁻-N and NO₃⁻-N removal rate of 8.8, 7.5 and 7.7 mg N L⁻¹ h⁻¹, respectively [90]. In addition, ammonium has also been demonstrated to be incorporated into biomass through assimilation pathways such as the glutamine synthetase-glutamate synthase (GS-GOGAT) system, thereby contributing to cellular nitrogen incorporation during the HNAD process [91]. Moreover, nitrogen balance analysis using isotope tracing method showed that 62.4% of the converted ammonium was assimilated into biomass in Corynebacterium glutamicum WN1, while 33.79%, 0.13%, and 0.04% were recovered as N₂-N, N₂O-N, and NH₂OH, respectively [21]. These findings indicate that assimilation contributes substantially to ammonium removal. The direct assimilation of ammonium may therefore partly explain the favorable growth and efficient substrate removal observed when ammonium is used as the sole nitrogen source. However, ammonium disappearance alone does not demonstrate heterotrophic nitrification. Distinguishing assimilatory uptake from other nitrogen transformations requires additional evidence, such as nitrogen-product measurements, isotope tracing, and nitrogen mass balances, depending on the experimental system.

NO₃⁻-N utilization may involve enzyme induction and sequential reduction. During complete aerobic denitrification, nitrate is reduced through nitrite, NO, and N₂O to N₂ by the corresponding reductases [92]. This process requires organic electron donors, making carbon availability and the C/N ratio important factors affecting nitrate removal [93]. An adaptation phase may occur when nitrate is supplied as the sole nitrogen source [94]. For example, Acinetobacter sp. HA2 exhibited a lag phase of approximately 8 h when switching from NH₄⁺-N to NO₃⁻-N as the sole nitrogen source, which was attributed to the induction of nitrate reductase synthesis [67].

NO₂⁻-N, an intermediate in both nitrification and denitrification, can exert concentration-dependent effects on HNAD activity. At relatively low concentrations, NO₂⁻-N can be rapidly transformed and may support denitrification [95,96]. At elevated concentrations, nitrite and its protonated form, free nitrous acid (FNA), may inhibit nitrogen-metabolizing enzyme activity and compromise cellular integrity, with their relative importance depending on pH and other cultivation conditions [97,98]. Nevertheless, some HNAD strains retain nitrogen-removal activity at elevated nitrite concentrations. For instance, Alcaligenes faecalis WT14 exhibited rapid growth and nearly complete removal of an initial NO₂⁻-N concentration of 100 mg N L⁻¹ within 24 h when nitrite was supplied as the sole nitrogen source at an initial pH of 7.0 and 30 °C [99]. Collectively, these findings indicate that some specific HNAD strains can efficiently utilize nitrite even at elevated concentrations; however, nitrite tolerance and inhibition should be evaluated in terms of both NO₂⁻-N concentration and pH-dependent FNA formation rather than nitrite concentration alone.

4. Environmental Responses of HNAD Bacteria

4.1. Effects of Carbon Source and C/N Ratio

Organic carbon availability is an important factor influencing nitrogen removal by HNAD bacteria, as organic carbon can provide energy, reducing power, and carbon skeletons for cell synthesis [8,100]. Under heterotrophic nitrification conditions, organic carbon supports biomass synthesis and cellular metabolism, but may not directly drive ammonia oxidation as an electron donor [34,45]. During aerobic denitrification, organic substrates can serve as electron donors for nitrate or nitrite reduction, support energy generation, and provide carbon for biomass synthesis [33,101,102]. Nevertheless, the extent of dependence on external organic carbon may vary among strains and experimental conditions [103,104]. Generally, carbon source type and C/N ratio jointly influence HNAD performance by modulating central carbon metabolism and the generation and partitioning of reducing equivalents, as schematically illustrated in Figure 2.

Figure 2. Conceptual framework illustrating carbon source and C/N ratio-regulated electron partitioning in HNAD bacteria.

4.1.1. Influence of Carbon Source on Heterotrophic Nitrification

Heterotrophic nitrification in HNAD bacteria is closely linked to intracellular carbon metabolism. The reducing power (e.g., nicotinamide adenine dinucleotide, NADH) and ATP, could be generated via glycolysis and the tricarboxylic acid (TCA) cycle during ammonium and carbon metabolism, which supports cellular growth and maintains redox balance [94]. The optimal carbon sources for different HNAD bacteria and the nitrogen removal rates were shown in Table 4.

Table 4. The optimal carbon source and C/N ratio for representative HNAD bacteria.

Strain	Carbon Source	Metabolic Entry Point	Optimal C/N Ratio	Removal Efficiency	Reference
Pseudomonas aeruginosa SNDPR-01	sodium citrate	TCA cycle intermediate	10	NH4+-N 87%	[26]
Acinetobacter ZHG-1	sodium citrate	TCA cycle intermediate	30	NH4+-N 97.1%	[105]
Paracoccus LYM	sodium lactate	Pyruvate precursor	15–30	NH4+-N 97.1%	[106]
Bacillus GS-5	sodium acetate	Short-chain fatty acid	7.5	NH4+-N 95.6%	[107]
Bacillus I6	glucose	Carbohydrate	8	NH4+-N 85%	[108]
Thauera sp. SND5	Sodium lactate	Pyruvate precursor	7.5	NH4+-N 99.2%	[109]
Cupriavidus S1	sodium pyruvate	Central metabolic intermediate	12–28	NH4+-N 99.7%	[30]
Bacillus LY	glucose	Carbohydrate	15	NH4+-N 100%	[110]
Zobellella sp. B307	sodium succinate	TCA cycle intermediate	5	NH4+-N 97.7%	[111]
Pseudomonas putida S3	sodium citrate	TCA cycle intermediate	6	NH4+-N 73.3%	[112]
Pseudomonas stutzeri AY4	glucose	Carbohydrate	6	NH4+-N 95.8%	[113]
	sodium succinate	TCA cycle intermediate		NO3−-N 91.4%
Chryseobacterium R31	glucose	Carbohydrate	10	NH4+-N 95.9%	[55]
	sodium succinate	TCA cycle intermediate

Existing research have shown that small organic acid salts generally outperform carbohydrates in promoting ammonium removal. For instance, in Pseudomonas sp. B-1, sodium succinate resulted in the highest ammonium removal efficiency (ARE) value of 57.5%, whereas glucose yielded only 16.1% [114]. Similar trends were observed in Acinetobacter tandoii MZ-5 [115] and Halomonas campislis ha3 [72], which exhibited enhanced ammonium removal when low-molecular-weight organic acids were used as carbon sources. This metabolic advantage can be attributed to the direct incorporation of substrates such as succinate and acetate into the TCA cycle, bypassing glycolysis and reducing metabolic burden, which ultimately leads to enhanced energy generation and stable redox conditions [23,44,116,117]. In contrast, carbohydrates typically undergo glycolysis and, in some cases, extracellular hydrolysis before entering central metabolism [23,118]. The additional enzyme-mediated reactions not only delay electron release but also divert carbon flux toward biomass synthesis [119]. Consequently, carbon source type regulates carbon flux distribution and electron availability within central metabolism, thereby influencing ammonium oxidation efficiency. Furthermore, different carbon sources might occupy distinct positions in cellular metabolic networks, influencing redox balance and enzyme regulation [120]. Accordingly, carbon sources that sustain continuous ATP generation and stabilize NADH/NAD⁺ balance are more favorable for efficient nitrogen transformation.

Overall, carbon sources can significantly influence HN through their effects on carbon flux, energy supply, and redox balance. In addition, low-molecular-weight organic acid salts usually favor ammonium removal despite of the different preferences among individual strains.

4.1.2. Influence of Carbon Source on Aerobic Denitrification

Unlike HN, AD is a reductive pathway that directly depends on intracellular reducing equivalents [96]. During aerobic denitrification, reducing equivalents generated through organic carbon metabolism can supply electrons for nitrate and nitrite reduction via electron-transfer components [15,101]. Therefore, carbon sources with high reducing power yield are expected to enhance denitrification efficiency.

Readily biodegradable carbon sources (e.g., acetate, citrate, succinate and other organic acid salts) have been reported to typically enhance aerobic denitrification rates by rapidly generating reducing power and promoting electron transfer to NO₃⁻/NO₂⁻ reductases [121]. For example, strain Acinetobacter Y1 exhibited significantly higher nitrogen removal efficiencies when utilizing sodium pyruvate, citrate, or acetate when compared with sucrose and glucose [122], which is similar to the strain Acinetobacter sp. ND7 [70]. In addition, carbon metabolism-derived ATP plays a crucial role in cellular energy metabolism and the activation of denitrification processes. In Acinetobacter johnsonii WGX-9, sodium acetate enhanced ATP synthesis and membrane transport activity, achieving an NO₃⁻ removal efficiencies around 86.6% at a rate of 0.2 mg N L⁻¹h⁻¹ [123]. A similar effect was also observed in Acinetobacter sp. TAC-1 [124]. It can therefore be inferred that carbon sources not only supply electrons but also influence enzyme synthesis through cellular energy status, thereby supporting efficient denitrification performance.

4.1.3. Effect of C/N Ratio on Nitrogen Removal Performance

Moderate-to-high C/N ratios (commonly 8–15) have been frequently reported to favor HNAD performance [31]. As shown in Table 4, increased carbon availability with increasing ratios generally enhances NADH and ATP production, thereby sustaining simultaneous oxygen respiration and aerobic denitrification [8,105]. Under carbon-limited conditions, the supply of reducing equivalents became insufficient to support both oxygen respiration and aerobic denitrification pathways, which usually resulted in incomplete denitrification or reduced ammonium oxidation efficiency.

Nevertheless, several HNAD strains maintain high nitrogen removal efficiencies even at low C/N ratios (≤3–4), including Janthinobacterium svalbardensis F19 (2) [125], Acinetobacter sp. Y16 (2) [126], and Acinetobacter haerbinensis HITLi7^T (3–4) [127]. These findings suggest that high carbon availability is not a universal prerequisite for efficient HNAD metabolism and that carbon requirements may vary with strain-specific differences in intracellular electron allocation and nitrogen transformation strategies [8,75]. In terms of electron and energy allocation, carbon-sensitive strains such as Acinetobacter junii ZHG-1 were strongly inhibited under carbon limitation due to decreased NADH/NAD⁺ ratios and reduced electron transport system activity (ETSA) [105]. In contrast, Bacillus thuringiensis WXN-23 maintained a high specific nitrogen removal rate under a relatively low C/N ratio (5.9), possibly by preferentially allocating available reducing equivalents toward nitrogen reduction rather than biomass synthesis [128]. The mechanistic basis underlying strain-specific electron allocation patterns and nitrogen transformation pathways in HNAD bacteria is further discussed in Section 5.2.

Overall, C/N dependency should not be interpreted solely as a preference for higher organic loading. It more fundamentally reflects the balance between electron generation and electron consumption within the cell [105]. When electron production exceeds respiratory demand, nitrogen removal is enhanced. While electron supply is limiting, oxygen respiration may outcompete denitrification [96]. In addition, strain-specific differences in this balance account for the observed variability in optimal C/N ratios. Understanding of the distinct metabolic patterns could provide new insights into the design of low-carbon nitrogen removal systems.

4.2. Effects of Environmental Factors

Environmental parameters can significantly influence enzyme activity and the allocation of intracellular reducing equivalents [129]. This section evaluated how key environmental factors, including temperature, pH, and DO, influence metabolic flux and electron transfer. The environmental characteristics and nitrogen removal performance of representative HNAD bacteria reported in previous studies were summarized in Table 5.

Table 5. Environmental characteristics and nitrogen removal performance of representative HNAD bacteria.

Strain	Nitrogen Source	Optimal Temperature (°C)	Optimal pH	DO Condition (rpm)	Nitrogen Removal Efficiency	Reference
Delftia tsuruhatensis SDU2	NH4+-N	30	7	150	NH4+-N 93.7%	[51]
Janthinobacterium sp. J1-1	NH4+-N; NO3−-N	8	7	150	NH4+-N 89.2%; NO3−-N 68.2%	[53]
Pseudomonas stutzeri strain AY4	NH4+-N; NO3−-N	30	7–8	180	NH4+-N 92.2%; NO3−-N 94.6%	[113]
Pseudomonas peli NR-5	NH4+-N; NO3−-N; NO2−-N	10	7	144	NH4+-N 97.3%; NO3−-N 95.3%; NO2−-N 87.8%	[130]
Paracoccus denitrificans AC-3	NH4+-N	35	6–10	150	NH4+-N > 99%	[131]
Pseudomonas tolaasii Y-11	NH4+-N; NO3−-N; NO2−-N	15	7.2	150	NH4+-N 93.6%; NO3−-N 93.5%; NO2−-N 81.9%	[27]
Acinetobacter baumannii AL-6	NH4+-N	30	7	120	NH4+-N 99.8%	[132]
Pseudomonas mosselii K17	NH4+-N	30	7	165	NH4+-N > 95%	[133]
Paracoccus denitrificans XW11	NH4+-N; NO3−-N	30	7.2	160	NH4+-N 100%; NO3−-N 90.5%	[129]
Acinetobacter ZHG-1	NH4+-N	30	9	150	NH4+-N 97.1%	[105]
Bacillus GS-5	NH4+-N	35	7.5	125	NH4+-N 95.6%	[107]
Chryseobacterium R31	NH4+-N	30	7	120	NH4+-N 99.7%	[55]
Vibrio diabolicus SF16	NH4+-N	30	7.2	120	NH4+-N 97.1%	[58]
Janthinobacterium sp. M-11	NH4+-N; NO3−-N; NO2−-N	2	7	150	NH4+-N 98%; NO3−-N 89%; NO2−-N 89%	[134]

4.2.1. Temperature

Temperature strongly regulates the metabolic activity and nitrogen removal performance of HNAD bacteria. The majority of documented HNAD strains exhibited a clear mesophilic preference, with optimal nitrogen removal typically occurring at 25–37 °C [135]. For example, Delftia tsuruhatensis SDU2 achieved the highest ammonia removal efficiency (93.7%) at 30 °C, whereas both cell growth and nitrogen removal decreased significantly at lower (20 °C) or higher (>35 °C) temperatures [51]. Similarly, Pseudomonas putida ZN1 showed negligible growth and denitrification activity at 10 °C or above 40 °C, whereas nitrogen removal efficiency exceeded 90% between 15 °C and 30 °C [88]. In general, nitrogen removal performance increased with temperature up to the optimum range (typically 25–37 °C), mainly owing to enhanced enzyme activity and microbial growth rates [136].

Although most HNAD bacteria exhibited mesophilic characteristics, growing evidence suggests that specific HNAD strains possess remarkable cold-tolerance capabilities and maintain efficient nitrogen removal performance. Cold-tolerant strains such as Janthinobacterium sp. J1-1 were able to achieve high removal of NH₄⁺-N (around 89.2%) and NO₃⁻-N (around 68.2%) at 8 °C [53]. In addition, psychrotolerant strains such as Bacillus simplex H-b and Priestia aryabhattai KX-3 have been reported to exhibit high ammonium removal efficiencies of 82.2% and 92.5%, respectively, even at 10 °C [101,137]. Strain-specific physiological and biochemical responses may contribute to cold tolerance in HNAD bacteria. In Acinetobacter calcoaceticus TY1, transcriptomic and biochemical analyses identified changes in fatty-acid composition, extracellular polysaccharide production, and antioxidant responses under cold stress [138]. These observations support a proposed role for membrane-related and oxidative-stress responses in maintaining cellular function at low temperatures. In Pseudomonas indoloxydans YY-1, increased ATP levels and extracellular polysaccharide content were reported following a temperature decrease, together with changes in the expression of genes involved in energy metabolism [139]. These measurements provide experimental evidence of cold-associated physiological responses. However, the contributions of individual responses to sustained nitrogen removal require further validation and should not be assumed to apply to all HNAD strains.

It is worth noting that thermophilic HNAD bacteria have also been identified. For instance, Anoxybacillus contaminans HA was reported to survive at temperatures up to 60 °C and exhibits optimal denitrification activity at 55 °C, with NH₄⁺-N and NO₃⁻-N removal efficiencies exceeding 70% [140]. However, the molecular mechanisms underlying temperature tolerance in HNAD bacteria remain insufficiently understood and require further investigation. The coexistence of psychrotolerant and thermophilic HNAD bacteria indicates substantial physiological plasticity in their nitrogen transformation pathways.

4.2.2. pH Value

pH is another critical environmental factor regulating the growth and nitrogen transformation activity of HNAD bacteria. Current studies have shown that HNAD bacteria can survive within a relatively wide pH range of 4–10 [141]. However, such viability does not necessarily imply comparable growth or nitrogen transformation activity throughout this range. In general, most strains exhibit optimal growth and nitrogen removal efficiency under neutral to slightly alkaline conditions [142,143]. For example, the ammonium removal efficiency of Pseudomonas mosselii K17 exceeded 85% within an initial pH range of 6–9 and reached approximately 95% at pH 7 [133].

Despite this general preference, several acid-tolerant HNAD bacteria have also been reported. For example, Acinetobacter sp. JR1 isolated from acidic pharmaceutical wastewater achieved a high ammonia removal efficiency of 97.9% at an initial pH of 4.5 [144]. In addition, Pseudomonas fragi EH-H1 was found to exhibit complete ammonium removal within 20 h at an initial pH of 6.2 [145]. These findings demonstrate that certain HNAD strains can retain ammonium removal activity under acidic initial conditions, although they do not by themselves establish sustained performance under fluctuating pH conditions.

The reported pH optima are generally condition- and strain-dependent, and multiple mechanisms may contribute to the pH response. Firstly, pH can influence membrane potential and proton gradients, thereby affecting ATP generation and nutrient transport [146,147]. In addition, extreme pH conditions can alter enzyme conformation and catalytic activity, thereby affecting nitrogen transformation activity [148]. Moreover, microorganisms may maintain intracellular pH homeostasis through proton transport systems and stress-response regulation [146]. These mechanisms may support pH tolerance, but do not necessarily preserve growth or nitrogen removal efficiency. Therefore, the observed pH optimum may be shaped by multiple interacting factors, including nitrogen speciation, membrane energetics, enzyme activity, substrate transport, and pH-dependent free-ammonia toxicity, with their relative contributions likely depending on the strain and cultivation conditions [149]. These mechanisms should be viewed as possible contributors to pH responses and tolerance, rather than as a complete explanation for the observed pH optimum of a given strain.

4.2.3. Dissolved Oxygen

DO is acknowledged to serve as a critical regulatory switch governing the HNAD process by directly mediating the cellular redox environment and intracellular electron distribution. Appropriate DO levels are essential to sustain the metabolic coupling between HN and AD; they provide sufficient oxygen for aerobic respiration and ammonia oxidation while ensuring that nitrogen oxides (NO₂⁻ and NO₃⁻) can still efficiently compete for electrons [150]. Driven by their rapid heterotrophic metabolism, many HNAD bacteria can quickly consume oxygen to create localized microaerobic niches within cellular aggregates, thereby enabling the concurrent operation of HN and AD pathways [151].

Changes in oxygen availability can affect nitrogen removal by altering the balance between aerobic respiration and nitrogen transformation. Low DO may limit oxygen-dependent ammonium oxidation and respiratory energy generation [152]. In contrast, high DO may favor oxygen respiration and reduce the availability of electrons for denitrification, particularly when the supply of reducing equivalents is limited [105,153]. The resulting effects on ammonium removal and denitrification depend on the strain, carbon availability, and cultivation conditions. DO should therefore be evaluated together with these factors when identifying suitable conditions for nitrogen removal.

Recent studies have underscored that the balance between oxygen utilization and electron allocation is maintained through diverse strain-specific physiological characteristics. For example, Pseudomonas sp. AND-42 exhibited the highest ammonium removal performance at a DO concentration of approximately 3.0 mg·L⁻¹, whereas higher DO levels (>8.0 mg·L⁻¹) resulted in a decline in NH4⁺-N removal efficiency, indicating the existence of a strain-specific oxygen optimum for coupled nitrogen conversion [154]. Similarly, Pseudomonas putida ZN1 achieved maximum ammonium removal (97.5%) at a shaking speed of 120 rpm corresponding to a DO concentration of 4.67 mg·L⁻¹, while either increasing or decreasing oxygen availability significantly reduced nitrogen removal performance [88]. In contrast, Pseudomonas stutzeri KTB exhibited a preference for relatively high oxygen availability and achieved complete nitrogen removal within 18 h at a shaking speed of 160 rpm [81]. Such tolerance to elevated oxygen availability may be associated with strong self-aggregation and rapid oxygen consumption, which have been reported to facilitate the formation of localized microaerobic niches and protect oxygen-sensitive denitrification enzymes in HNAD bacteria [155,156]. Overall, DO regulates both HN and AD processes by influencing oxygen availability and electron allocation. Effective DO control should therefore accommodate strain-specific physiological characteristics to support aerobic respiration and ammonia oxidation as well as maintaining sufficient electron availability for denitrification.

4.2.4. Interactive Effects of the Environmental Factors

The nitrogen source serves as a critical nexus linking carbon availability, oxygen supply, and the effects of pH and temperature. Assimilation of nitrate or nitrite imposes a higher reducing-equivalent demand than ammonium assimilation [157], thereby rendering the C/N response partly contingent on the nitrogen substrate. Given that reducing equivalents derived from carbon metabolism are also utilized by aerobic respiration, an elevated DO level may impose a constraint on denitrification when carbon availability is limiting [105,153]. The nitrogen source further governs the relevance of pH-dependent speciation: elevated pH and temperature raise the free ammonia fraction in ammonium-containing media, whereas reduced pH increases the free nitrous acid fraction in the presence of nitrite [98,158]. Thus, nitrogen removal reflects both the availability of electrons for the relevant pathways and the chemical forms of nitrogen under the cultivation conditions. Operating optima should therefore be evaluated for specific combinations of nitrogen source, C/N, DO, pH, and temperature.

4.3. Effects of Extreme Conditions on HNAD Bacteria

Extreme environmental conditions, particularly high salinity and heavy metal stress, can significantly affect the nitrogen removal performance of HNAD bacteria. Salinity imposes osmotic stress that disrupts cellular metabolism, whereas heavy metals may inhibit key enzymes involved in nitrogen transformation. Nevertheless, several HNAD strains have been reported to sustain efficient nitrogen removal activity under high salinity and heavy metal stress conditions as summarized in Table 6.

Table 6. Nitrogen removal performance of HNAD bacteria under high salinity and heavy metal stress.

Stress Condition	Strain	Stress Level	Nitrogen Removal Efficiency	Reference
High salinity	Zobellella endophytica W14	5% NaCl	NH4+-N 100%; NO3−-N 98.9%; NO2−-N 99.6%	[43]
	Halomonas sp. 5505	8% NaCl	NH4+-N 100%; NO3−-N 94.1%; NO2−-N 74.4%	[85]
	Acinetobacter sp. JR1	20 g·L−1 NaCl	NH4+-N 96.0%	[144]
	Serratia marcescens CL1502	1–5% NaCl	NH4+-N > 90.0%	[159]
	Zobellella sp. B307	75 g·L−1 NaCl	NH4+-N 97.7%; NO3−-N 94.4%	[111]
	Halomonas sp. B01	60 g·L−1 NaCl	NH4+-N 98.8%	[160]
	Bacillus N31	30–40‰ NaCl	NH4+-N > 80.17%	[71]
	Pseudomonas aeruginosa P-1	40 g·L−1 NaCl	NH4+-N 100%	[161]
heavy metals	Pseudomonas putida LX1	25.0 mg·L−1 Cr6+	NH4+-N 83.4%	[36]
	Aeromonas sp. HN-02	8.0 mg·L−1 Zn2+; 0.5 mg·L−1 Cu2+ + Zn2+	NH4+-N 94.6%; NH4+-N 46.4%	[46]
	Acinetobacter sp. JR1	20.0 mg·L−1 Mn2+	NH4+-N 90.5%	[144]
	Pseudomonas putida NP5	50.0 mg·L−1 Cr6+	NH4+-N > 40%	[17]
	Pseudomonas putida ZN1	60.0 mg·L−1 Cr6+; 20.0 mg·L−1–80.0 mg·L−1 Ni2+	NH4+-N 94.9%; NH4+-N 71.3–31.5%	[88]
	Pseudomonas aeruginosa P-1	30 mg·L−1–50 mg·L−1 Cu2+; 50 mg·L−1 Cr6+	NH4+-N 100–52.18%; NH4+-N > 80%	[161]

4.3.1. Effect of High Salt on HNAD Bacteria

High-salinity wastewater, typically defined as wastewater containing NaCl concentrations exceeding 3% (w/v) [162], is widely generated from industrial processes such as petroleum exploitation, marine aquaculture, food processing, pharmaceutical manufacturing, and landfill leachate concentration processes [163,164]. Elevated salt concentrations could impose osmotic stress on microbial cells, which can lead to cellular dehydration, disruption of membrane integrity, and reduced enzyme activity [165]. For HNAD bacteria, salinity stress significantly affected both HN and AD processes through alterations in cellular metabolism and osmotic balance [166]. Under high salinity conditions, the increase in extracellular osmotic pressure can cause water efflux through cell membrane, resulting in reduced metabolic activity and slower growth rates [167]. In addition, salinity might suppress the activity of key enzymes such as ammonia monooxygenase (AMO) and nitrate reductase (NR), thereby limiting nitrogen conversion efficiency [166].

Despite the inhibitory effects, a number of HNAD bacteria have demonstrated remarkable tolerance potentials under saline environments. Several salt-tolerant HNAD strains isolated from marine sediments, saline soils, and aquaculture systems with remarkable nitrogen removal capacity have been identified in recent years [8,89], mainly including genus Acinetobacter, Halomonas, Zobellella, Bacillus, Pseudomonas, Alcaligenes, and Flavobacterium (Table 6). For instance, Zobellella endophytica W14, isolated from hypersaline wastewater by Hu et al., was able to grow and efficiently remove ammonium in environments with salinity ranging from 0 to 11% (w/v) [43]. Similarly, Yao et al. reported that Bacillus cereus X7 achieved an ammonium removal efficiency exceeding 97.6% at NaCl concentrations ranging from 0 to 40·g L⁻¹ [168]. Similar salt-tolerant characteristics have also been observed in HNAD-based microbial communities operating under saline conditions. Zhang et al. have developed a HNAD bacteria-functionalized biocathode for treating high-salinity ammonium-rich wastewater and reported the TN removal efficiency of 92.5% with stable operation over 350 days [169]. Microbial community analysis revealed the enrichment of salt-tolerant electroactive HNAD bacteria, particularly the genus Vitellibacter, which suggested that the coexistence of halotolerance and extracellular electron transfer capability contributed to the sustained nitrogen removal performance under saline conditions [169].

Several protective mechanisms have been proposed to explain salt tolerance in HNAD bacteria. Under saline conditions, the accumulation of compatible solutes, such as ectoine, trehalose, glycine betaine, and proline, may support osmotic balance and help stabilize cellular structures and enzymes in salt-tolerant HNAD strains [8,170]. Hu et al. obtained Halomonas salifodinae cultures with different salt tolerances after three cultivation cycles at different salinities [171], indicating that tolerance depended on cultivation history. Metabolite analysis suggested that compatible-solute accumulation contributed to this acclimation response [171]. High salinity may also induce the formation of reactive oxygen species (ROS); therefore, enhanced antioxidant defense systems may contribute to alleviating oxidative damage and maintaining cellular metabolism [160]. In addition, regulation of key nitrogen metabolism enzymes, including AMO and NR, may help certain strains sustain active HNAD pathways under saline conditions [172]. For instance, intracellular ectoine concentrations in Halomonas sp. B01 increased with rising NaCl concentrations, while superoxide dismutase (SOD) activity increased significantly at 120 g L⁻¹ NaCl [160]. These observations suggest that osmotic regulation and antioxidant responses may contribute to salt tolerance in this strain.

4.3.2. Effect of Heavy Metals on HNAD Bacteria

Heavy metal contamination is another important environmental stressor. Heavy metals such as Cu, Zn, Cr, and Ni are commonly present in industrial effluents generated by metallurgy, electroplating, printing and dyeing, and tanning industries [88]. These metals may bind to cellular proteins and enzymes, disrupt membrane permeability, and interfere with electron transport chains, thereby affecting microbial metabolism [173]. In addition, heavy metals may directly inhibit key enzyme activities involved in nitrogen transformation. Enzymes responsible for aerobic denitrification, including NR and nitrite reductase, can be inhibited through metal binding to active sites or through the generation of reactive oxygen species. As a result, the overall nitrogen removal efficiency may decline under elevated metal concentrations.

Nevertheless, certain HNAD bacteria have demonstrated notable tolerance to heavy metal stress and can maintain efficient nitrogen removal performance in metal-contaminated environments, as summarized in Table 6. For example, Pseudomonas aeruginosa P-1 was reported to achieve an ammonia removal efficiency above 80% at Cr⁶⁺ concentration of 50.0 mg·L⁻¹, with negligible nitrite accumulation [161]. Similarly, Pseudomonas putida ZN1 was observed to obtain average NH₄⁺-N removal efficiency of 94.9% and 92.1% under 60 mg·L⁻¹ Cr⁶⁺ and 8.0 mg·L⁻¹ Cu²⁺ stress conditions, respectively [88].

Several protective mechanisms have been proposed to explain heavy metal tolerance in HNAD bacteria. Extracellular polymeric substances (EPS) can adsorb and immobilize metal ions, thereby reducing their direct toxicity to cells [36]. In addition, metal resistance systems, detoxification pathways, and antioxidant defenses may help alleviate metal-induced stress and maintain cellular homeostasis [174]. However, their contribution to nitrogen removal during stress exposure and recovery remains to be further elucidated for individual strains. Moreover, current insights into salinity and metal tolerance may be insufficient to fully determine the rate and extent to which nitrogen transformation activity recovers after stress relief, or whether prior exposure alters subsequent stress responses. These uncertainties may currently constrain predictions of treatment stability under fluctuating conditions.

5. Nitrogen Metabolic Mechanism of HNAD Bacteria

5.1. Nitrogen Transformation Pathways

Enzymes implicated in reported or proposed HNAD pathways mainly include AMO, hydroxylamine oxidase (HAO), nitrite oxidoreductase (NXR), nitrate reductases, NirK/NirS, Nor, and NosZ. However, their occurrence and functional characterization vary among strains. Direct biochemical evidence, including protein purification, has been obtained for AMO and hydroxylamine oxidase in selected heterotrophic nitrifiers. For example, an active ammonia monooxygenase was purified from Paracoccus denitrificans [175], whereas a hydroxylamine oxidase distinct from that of autotrophic nitrifiers was purified from Thiosphaera pantotropha [13]. The biochemical basis of ammonia and hydroxylamine oxidation is not yet fully understood across HNAD bacteria, with enzyme assignments in many strains based on genomic annotation or indirect physiological evidence [10]. Similarly, nitrate reduction may involve membrane-bound nitrate reductase (Nar), periplasmic nitrate reductase (Nap), or assimilatory nitrate reductase (Nas), depending on the organism and physiological context. Nas is generally considered to participate in nitrogen assimilation and is therefore distinguished from nitrate reductases associated with denitrification [157].

During nitrogen metabolism, nitrogen may occur as NH₄⁺, NO₂⁻, NO₃⁻, NH₂OH, NO, N₂O and N₂ [8]. Reported HNAD pathways can be broadly grouped into two representative categories. One involves coupled nitrification and denitrification through NO₂⁻/NO₃⁻ intermediates [34,126]. The other comprises proposed NH₂OH-mediated routes to gaseous nitrogen products without NO₃⁻ or NO₂⁻ as obligatory intermediates [176,177,178]. These categories reflect the transformation patterns currently reported, although the underlying reactions and enzymes have not yet been equally well resolved across strains. In addition to these pathways, nitrogen assimilation contributes to ammonium removal through incorporation into biomass. Reported or proposed HNAD pathways, representative strains, and their ammonium removal efficiencies are summarized in Table 7.

Table 7. Reported or proposed nitrogen transformation pathways in bacteria described as HNAD strains and their ammonium removal performance.

Pathways	Reported or Proposed Nitrogen Transformation Pathway	Strain	NH4+-N Removal Efficiency (%)	Reference
NO3−/NO2−-mediated pathways	NH4+ → NH2OH → NO2− → NO3− → NO2−→ NO → N2O→N2	Pseudomonas stutzeri KTB	NH4+-N 100%	[81]
		Paracoccus denitrificans HY-1	NH4+-N 98.1%	[120]
		Providencia rettgeri YL	NH4+-N 80–97%	[47]
		Ochrobactrum anthropic LJ81	NH4+-N > 80%	[54]
		Delftia sp. B7	NH4+-N 98.7%	[179]
		Pseudomonas sp. JQ-H3	NH4+-N 95.4%	[180]
	NH4+ → NH2OH → NO2− → NO3− → NO2−→ NO → N2O	Pseudomonas sp. ZHL02	NH4+-N 99.1%	[93]
	NH4+ ↔glutamine→ NH2OH → NO2− → NO3− → NO2−→ NO → N2O→N2	Zobellella sp. An-6	NH4+-N 96.1%	[181]
	NH4+ → NH2OH → NO2− → NO3− → NO2− → N2O→N2	Acinetobacter junii YB	NH4+-N 99.9%	[176]
	NH4+ → NH2OH → NO2− → NO3− → NO2−→N2	Pseudomonas fluorescens wsw-1001	NH4+-N 71.2%	[182]
		Paracoccus versutus LYM	NH4+-N 75.4%	[183]
		Lysinibacillus fusiformis B301	NH4+-N 83.3%	[84]
	NH4+ → NH2OH → NO2− → NO3−→ N2O→N2	Bacillus subtilis A1	NH4+-N 58.4%	[184]
		Rhodococcus sp. CPZ24	NH4+-N 85%	[57]
		Klebsiella pneumoniae CF-S9	NH4+-N 85%	[80]
	NH4+ → NH2OH → NO2− → N2O→N2	Agrobacterium sp. LAD9	NH4+-N 90.1%	[23]
		Alcaligenes ammonioxydans sp. HO-1	NH4+-N 84%	[41]
	NH4+ → NH2OH → NO2− → NO→ N2O→N2	Pseudomonas stutzeri YG-24	NH4+-N 88.1%	[90]
NH2OH-mediated pathways	NH4+ → NH2OH → NO → N2O→N2	Alcaligenes faecalis TF-1	NH4+-N 94.6%	[74]
	NH4+ → NH2OH → N2O→N2	Bacillus methylotrophicus L7	NH4+-N 91%	[66]
		Acinetobacter calcoaceticus HNR	NH4+-N 92.3%	[178]
		Photobacterium sp. NNA4	NH4+-N > 99.9%	[135]
		Alcaligenes faecalis No. 4	NH4+-N > 99.9%	[185]
		Alcaligenes faecalis NR	NH4+-N 66.7–78.3%	[186]
	NH4+ → NH2OH →N2	Corynebacterium glutamicum WN1	NH4+-N > 99.9%	[21]
Other novel pathways	NH4+ → NO → N2O	Pseudomonas putida Y-9	NH4+-N 99.41%	[187]
	NH4+ → N2O→N2	Pseudomonas taiwanensis sp. J488	NH4+-N > 99.9%	[188]

5.1.1. NO₃⁻/NO₂⁻-Mediated Pathways

Wehrfritz et al. proposed a model linking heterotrophic nitrification to aerobic denitrification through shared electron-transfer components in 1993 [13]. A proposed NO₃⁻/NO₂⁻ -mediated pathway is illustrated in Figure 3, in which ammonium is oxidized to nitrite via hydroxylamine, with subsequent oxidation to nitrate in some strains. AMO and HAO have been reported in T. pantotropha [45], while NXR activity has been reported in Glutamicibacter arilaitensis EM-H8 [189]. These observations provide biochemical context for the model rather than verification of the complete pathway within a single strain. In the denitrification module, NO₃⁻ is reduced to NO₂⁻ by a dissimilatory nitrate reductase, such as Nap or Nar, depending on the strain and environmental conditions (Equation (1)). NO-forming NirK/NirS subsequently reduce NO₂⁻ to NO (Equation (2)), Nor reduces NO to N₂O (Equation (3)), and NosZ catalyzes the reduction of N₂O to N₂ (Equation (4)) [190,191]. When all these denitrification steps occur in coupling with the upstream nitrification steps, the overall nitrogen removal sequence can be represented as NH₄⁺ → NH₂OH → NO₂⁻ → NO₃⁻ → NO₂⁻→ NO → N₂O→N₂. This integrated sequence provides a framework for the reported HNAD pathways, although the specific enzymes and intermediates may vary among strains.

Figure 3. Schematic overview of reported and proposed nitrogen transformation pathways in HNAD bacteria.

NO₃⁻/NO₂⁻ -mediated pathways have been reported or proposed for several HNAD strains exhibiting high ammonia removal efficiencies. For example, Delftia sp. B7 achieved an NH₄⁺-N removal efficiency of 98.7 ± 0.2%, and a removal rate of 4.1 ± 0.04 mg N·L⁻¹h⁻¹ [179]. Nitrogen transformation experiments and genomic analyses supported a proposed NO₃⁻/NO₂⁻-mediated pathway in this strain. However, canonical amoCBA and hao genes were not identified in the available genome annotation; consequently, the genetic and enzymatic basis of the initial ammonia oxidation steps remains to be further elucidated [179].

2NO₃⁻ + 4H⁺ + 4e⁻ → 2NO₂⁻ + 2H₂O

(1)

2NO₂⁻ + 4H⁺ + 2e⁻ → 2NO + 2H₂O

(2)

2NO + 2H⁺ + 2e⁻ → N₂O + H₂O

(3)

N₂O + 2H⁺ + 2e⁻ → N₂ + H₂O

(4)

In addition, variations of the NO₃⁻/NO₂⁻-mediated pathways have been reported, differing mainly in the involvement of NO₃⁻ and the intermediates included in the proposed sequences (Table 7). These variants can be generally grouped into (i) pathways involving NO₃⁻ formation before denitrification, (ii) pathways in which NO₂⁻ enters denitrification without prior oxidation to NO₃⁻, and (iii) abbreviated reported sequences in which intermediates such as NO or N₂O are not explicitly represented. Importantly, the omission or non-detection of an intermediate might not be taken as evidence that the corresponding biochemical step is bypassed. The observed diversity in nitrogen transformation pathways suggests that nitrogen transformation varies among HNAD strains [8].

5.1.2. Proposed NH₂OH-Associated Pathways

In the proposed NH₂OH-associated pathways, NH₂OH serves as a putative intermediate linking ammonium oxidation to gaseous nitrogen formation. Several NH₂OH-associated routes involving NO and/or N₂O have been reported and are summarized in Figure 3, with representative strains listed in Table 7. The proposed routes mainly include NH₄⁺ → NH₂OH → NO → N₂O →N_2, NH₄⁺ → NH₂OH → N₂O→N₂ and NH₄⁺ → NH₂OH → N₂, differing in the intermediates proposed between NH₂OH and N₂.

For Alcaligenes faecalis TF-1, a route involving NO and N₂O as intermediates was proposed based on PCR detection of amoA, hao, norB, and nosZ, together with substrate-utilization experiments, which exhibited ammonium removal but no detectable aerobic denitrification when nitrate or nitrite was supplied as the sole nitrogen source [74]. These results support a proposed route but do not, by themselves, establish NH₂OH, NO, or N₂O as intermediates or demonstrate the activities of the corresponding enzymes. In Photobacterium sp. NNA4, NH₂OH consumption accompanied by N₂O production was observed in cell-suspension experiments with uninoculated and heat-inactivated controls, and HAO activity was detected in cell-free extracts. These findings supported the proposed conversion of NH₂OH to N₂O. Separate experiments with ¹⁵N-labeled nitrate and nitrite further demonstrated aerobic production of N₂O and N₂ [135], but did not directly establish the complete route from ammonium through NH₂OH to gaseous nitrogen. For Corynebacterium glutamicum WN1, the NH₄⁺ → NH₂OH → N₂ route was proposed by combining NH₂OH measurements, ¹⁵NH₄⁺ tracing, gaseous-product analysis, and genomic and transcriptomic analyses [21]. The direct NH₄⁺ → NH₂OH → N₂ route is proposed but not yet experimentally verified. Trace NH₂OH and ammonium-derived ¹⁵N₂ were detected, and genes annotated as dnfB and dnfC were upregulated following ammonium addition [21]. However, the absence of detectable NO₂⁻, NO₃⁻, or N₂O does not rule out their involvement in the proposed routes, as rapid formation and consumption may maintain their concentrations below analytical detection limits.

The proposed NH₂OH-associated pathways may provide alternative routes for nitrogen removal under aerobic conditions. By limiting accumulation of potentially toxic intermediates such as NO₂⁻ and NO, these pathways may help alleviate cellular stress [93]. Overall, the proposed NH₂OH-associated pathways highlight the potential diversity of nitrogen metabolism in HNAD bacteria.

5.1.3. Nitrogen Assimilation

While the previous sections primarily focus on dissimilatory pathways that convert inorganic nitrogen into gaseous products, HNAD bacteria can also assimilate inorganic nitrogen into cellular biomass [19,192]. The assimilation process represents an important nitrogen retention mechanism in HNAD bacteria. Therefore, nitrogen transformation in HNAD bacteria is expected to be governed by the balance between assimilatory nitrogen uptake and dissimilatory nitrogen loss. In most bacteria, ammonium assimilation primarily occurs via the glutamine synthetase-glutamate synthase (GS-GOGAT) pathway [193]. The relevant enzymes involved in nitrogen assimilation include glutamine synthase (GS), glutamate synthase (GOGAT) and glutamate dehydrogenase (GDH) [194,195]. In this process, ammonium is first incorporated into glutamine via glutamine synthetase and subsequently converted into glutamate through glutamate synthase. Glutamate then serves as a central nitrogen donor for the biosynthesis of amino acids, nucleotides, and other nitrogen-containing cellular components as illustrated in Figure 3 [91]. Through the assimilatory pathway, inorganic nitrogen can be effectively incorporated into microbial biomass, thereby supporting cellular growth and metabolic activity.

The contribution of nitrogen assimilation varies from different HNAD strains and the representative strains and their reported assimilation contributions are summarized in Table 8. For instance, nitrogen assimilation was the dominant nitrogen removal pathway in Klebsiella sp. under aerobic condition [196], as well as Pseudomonas putida NP5 can assimilate up to 61.8% of ammonia nitrogen [17]. In contrast, only 24.0% of nitrogen is removed via assimilation in Rhodococcus sp. CPZ24 [57]. Therefore, the balance between nitrogen assimilation and dissimilatory nitrogen removal is highly strain-dependent, reflecting the allocation of nitrogen metabolic flux between biosynthetic demand and energy-conserving dissimilatory processes.

Table 8. Contribution of assimilatory nitrogen metabolism in reported HNAD bacteria.

Strain	Nitrogen Source	Assimilation Contribution (%)	Dominant Nitrogen Pathway	Reference
Bacillus sp. LY	NH4+-N	40.5%	Multiple pathways	[110]
Providencia rettgeri strain YL	NH4+-N	49.7%	Multiple pathways	[47]
Agrobacterium sp. LAD9	NH4+-N	40.8%	Multiple pathways	[23]
Pseudomonas putida NP5	NH4+-N	61.8%	Assimilation-dominated	[17]
Acinetobacter haemolyticus JS-1	NH4+-N; NO3−-N	74.5%; 85.9%	Assimilation-dominated	[197]
Rhodococcus sp. CPZ24	NH4+-N	24%	Multiple pathways	[57]
Enterobacter asburiae GS2	NH4+-N	63.7%	Assimilation-dominated	[198]
Halomonas sp. B01	NH4+-N;	39.4%	Multiple pathways	[160]
Halophilic Vibrio diabolicus SF16	NH4+-N	35.8%	Multiple pathways	[58]
Alcaligenes faecalis C16	NH4+-N	28–45%	Multiple pathways	[199]
Acinetobacter calcoaceticus HNR	NH4+-N	52.1%	Assimilation-dominated	[178]
Corynebacterium glutamicum WN1	NH4+-N	62.4%	Assimilation-dominated	[21]
Ochrobactrum anthropi HND19	NH4+-N; NO3−-N; NH4+-N + NO3−-N	37.9%; 26.4%; 27.9%	Multiple pathways	[200]
Ochrobactrum anthropic LJ81	NH4+-N; NO3−-N; NO2−-N	9.5%; 10.6%; 11.1%	Multiple pathways	[54]
Paracoccus denitrificans HY-1	NH4+-N	80.7%	Assimilation-dominated	[120]

The relative contribution of assimilatory nitrogen metabolism in HNAD bacteria is significantly influenced by environmental conditions, particularly the C/N ratio and nitrogen sources. Several studies have suggested that elevated C/N ratios could promote nitrogen assimilation and biomass formation under carbon-sufficient conditions, thereby increasing nitrogen retention within microbial cells [197]. For example, at a C/N ratio of 20, the marine bacterium Halomonas sp. DN3 exhibited dominant nitrogen assimilation, with 75.0–85.8% of the removed nitrogen assimilated into cellular biomass [201]. From a metabolic perspective, the assimilatory shift under high C/N ratios is associated with increased availability of carbon skeletons, reducing equivalents (NADH/NADPH), and ATP generated through enhanced carbon oxidation [105]. These metabolic resources might facilitate ammonium incorporation through key assimilation pathways, including the GDH and GS-GOGAT routes, thereby promoting intracellular nitrogen retention rather than extracellular nitrogen transformation [202].

The type of nitrogen source is a critical determinant of assimilatory nitrogen metabolism in HNAD bacteria, owing to differences in energetic requirements and enzymatic regulation. For example, for Halomonas sp. B01, nitrogen assimilation accounted for 39.4%, 59.4%, and 56.2% of nitrogen removal when NH₄⁺-N, NO₃⁻-N, NO₂⁻-N were used as the sole nitrogen sources, respectively [160]. Such variation reflects not only strain-specific metabolic characteristics but also the different energetic costs associated with assimilation of reduced and oxidized nitrogen compounds [203]. NH₄⁺ is generally considered the most energetically favorable nitrogen source because it can be directly incorporated into cellular metabolites without prior reduction [201]. In contrast, the assimilation of NO₃⁻ and NO₂⁻ requires additional reduction steps mediated by assimilatory nitrate and nitrite reductases (Nas systems), consuming reducing equivalents and metabolic energy [204]. Under carbon-rich conditions, sufficient electron and energy availability may facilitate these reductive assimilation processes and increase nitrogen retention in biomass.

Finally, assimilatory nitrogen metabolism may enhance the adaptability of HNAD bacteria to environments with elevated ammonium availability. Compared with dissimilatory pathways, ammonium assimilation involves fewer reaction steps, which may facilitate rapid nitrogen incorporation into cellular biomass [40].

5.2. The Proposed Nitrogen Metabolic Pathways

The ability of HNAD bacteria to couple HN and AD under aerobic conditions is closely associated with the coordination of carbon metabolism, electron transport processes, and nitrogen transformation pathways [8]. A conceptual framework of this integrated metabolic network is presented in Figure 4, illustrating the proposed distribution of electron flux between oxygen respiration and nitrogen oxide reduction, as well as its linkage to carbon metabolism and enzymatic nitrogen conversions.

Figure 4. Proposed mechanistic model of nitrogen metabolism in HNAD bacteria.

During carbon metabolism, organic substrates are oxidized via glycolysis and the tricarboxylic acid (TCA) cycle, generating ATP through aerobic respiration and reducing equivalents (e.g., NADH and FADH₂) that support cellular metabolism and nitrogen transformation [119]. The intracellular NADH/NAD⁺ ratio reflects cellular redox status and may be associated with changes in respiratory and nitrogen transformation activities [205]. The reducing equivalents subsequently supply electrons (e⁻) to the respiratory ETC, which may link carbon oxidation with nitrogen transformation [206]. Within the ETC, electrons derived from NADH can be partitioned among multiple terminal electron acceptors. Under aerobic conditions, O₂ typically functions as the primary electron acceptor [129], whereas nitrogen oxides such as NO₃⁻ and NO₂⁻ can serve as alternative acceptors in HNAD bacteria [95]. Consequently, reducing equivalents generated through organic carbon oxidation may also support denitrification in the presence of oxygen [93]. The proposed allocation of electrons between oxygen respiration and nitrogen oxide reduction provides a possible explanation for aerobic denitrification (Figure 4). Within this framework, carbon metabolism and electron transport may help coordinate the oxidative reactions of heterotrophic nitrification with the reductive reactions of denitrification.

As described in Section 5.1, enzymes associated with HNAD pathways have been reported in individual strains, although their functional characterization varies. For example, nitrogen balance analysis and enzyme assays in Paracoccus versutus LYM supported a proposed pathway linking heterotrophic nitrification with aerobic denitrification [183]. Heterotrophic carbon metabolism provides reducing equivalents that may support both respiratory oxygen reduction and nitrogen oxide reduction. The proposed allocation of reducing equivalents, together with the potential coexistence of enzymes involved in nitrification and denitrification, may support concurrent oxidative and reductive nitrogen transformations under aerobic conditions [105].

6. Challenges and Future Perspectives

Although environmental factors, including DO, carbon source type, temperature, salinity, and heavy metal stress, are well recognized as key regulators of HNAD activity, how these factors reshape intracellular redox balance, enzyme activity, and pathway selection remains insufficiently understood. In particular, identifying the conditions that drive shifts in the relative contributions of nitrification, denitrification, and assimilation pathways is essential for developing control strategies to improve system performance and maintain stable operation. The extent and timescale of activity recovery after salinity returns to baseline or inhibitory metals are removed also remain insufficiently characterized in HNAD isolates. Controlled exposure-recovery and re-exposure experiments are needed to assess recovery kinetics, prior exposure effects, and potential irreversible activity loss.

In addition, nitrogen transformation in HNAD bacteria is intimately coupled with carbon metabolism and electron transport. Nevertheless, how electrons are allocated among biomass synthesis, respiratory energy conservation, and nitrogen conversion pathways remains poorly quantified, especially under variable environmental conditions. To address this knowledge gap, future studies should couple measurements of substrate utilization kinetics, carbon consumption, oxygen uptake, and key enzyme activities with nitrogen mass balances that encompass dissolved, biomass-associated, and gaseous nitrogen. Such integrated analyses would distinguish assimilatory nitrogen retention from gaseous nitrogen production and help constrain electron allocation among competing metabolic processes.

Furthermore, although the major nitrogen conversion pathways in HNAD systems—encompassing ammonia oxidation, nitrate/nitrite reduction, and assimilation—have been postulated, their enzymatic underpinnings and relative contributions under aerobic conditions remain poorly characterized. The observation of ammonia oxidation activity in certain strains lacking canonical amo and hao genes suggests the possible involvement of alternative enzymatic systems or unidentified routes. Hence, future studies should prioritize the elucidation of key functional enzymes and intermediate metabolites, especially those involved in the hydroxylamine-mediated pathways. The integration of enzyme activity assays with time-resolved nitrogen transformation rates would facilitate linking biochemical functions to in situ pathway activities under variable environmental conditions.

Finally, although HNAD bacteria have been applied in laboratory-scale treatment of real wastewater, including livestock-farm and aquaculture wastewater using Pseudomonas stutzeri AY4 and Pseudomonas putida 4-HYK, respectively [90,207], these demonstrations have yet to establish reliable full-scale performance. Wider application requires addressing carbon and aeration demands, population stability under fluctuating conditions, and potential nitrite accumulation or N₂O emissions. For example, although HNAD can simplify treatment by integrating nitrogen and COD removal in a single aerobic system compared with conventional nitrification-denitrification [90], realizing this advantage requires optimization of C/N ratio and DO to sustain nitrogen removal rates without excessive carbon inputs or aeration [208]. In addition, key functional enzymes in HNAD, such as nitrate reductase, are oxygen-sensitive, and DO levels strongly affect nitrogen removal and N₂O emissions [209]. Compared with anaerobic denitrification processes, HNAD exhibits faster growth and greater environmental adaptability, particularly in high-organic-strength wastewater; however, competition for carbon sources among functional microbial groups and N₂O release have yet to be fully resolved [210]. Furthermore, although the kinetics of pure HNAD strains can be described by Monod or Haldane models, the kinetic parameters of HNAD and the microbial community stability in pilot-scale or full-scale treatment systems still need to be further addressed, particularly the specific nitrogen removal rate and yield coefficients of HNAD bacteria. Overall, HNAD shows considerable promise for treating wastewater with a high C/N ratio in single-reactor systems under energy-efficient operation. Nevertheless, further studies are required to address strain screening, process control, kinetic modeling, N₂O emission mitigation, and the long-term stability of both strains and microbial communities.

7. Conclusions

HNAD bacteria constitute a diverse and functionally distinct microbial guild capable of simultaneous HN and AD, with potential for biological nitrogen removal in wastewater treatment. This review systematically summarizes the current understanding of HNAD bacteria from the perspectives of distribution, physiological characteristics, environmental responses, nitrogen transformation pathways, and metabolic mechanisms. Bacterial isolates reported to exhibit HNAD activity have been successfully isolated from diverse natural and engineered environments, exhibiting considerable physiological versatility across strains. The nitrogen removal performance of HNAD bacteria is governed by the interplay between nitrogen sources, carbon sources, C/N ratio, and environmental conditions (e.g., temperature, pH, dissolved oxygen, salinity, and heavy metals). Reported nitrogen transformation routes include NO₃⁻/NO₂⁻-mediated pathways, NH₂OH-associated pathways, and nitrogen assimilation. Assimilation can contribute substantially to inorganic nitrogen removal through biomass incorporation, and the fate of nitrogen is governed by the balance between dissimilatory transformation and assimilation. Collectively, the physiological versatility and metabolic flexibility of HNAD bacteria highlight their potential for application in sustainable biological nitrogen removal systems. Future research should quantify assimilatory and dissimilatory contributions through comprehensive nitrogen mass balances, identify key enzymes or genes involved, elucidate electron allocation mechanisms, and validate long-term treatment performance under conditions representative of real wastewater.