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Article

Adopting Biochar as Immobilization Support for Hyper Ammonia-Producing Bacteria Proliferation

by
Christiana Bitrus
1,
Ademola Hammed
1,2,*,
Tawakalt Ayodele
1,
Kudirat Alarape
1,
Niloy Chandra Sarker
2,
Clairmont Clementson
2 and
Ewumbua Monono
2
1
Environmental and Conservation Science, North Dakota State University, Fargo, ND 58102, USA
2
Agricultural and Biosystems Engineering, North Dakota State University, Fargo, ND 58102, USA
*
Author to whom correspondence should be addressed.
Appl. Microbiol. 2025, 5(4), 111; https://doi.org/10.3390/applmicrobiol5040111
Submission received: 18 August 2025 / Revised: 9 October 2025 / Accepted: 10 October 2025 / Published: 14 October 2025
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Abstract

The many uses of biochar extend to microbial enhancement in fermentation processes because it acts as a catalyst and a support medium in agricultural industries, particularly for biofertilizer production. This study explores how three key biochar parameters, concentration (0.05–0.25% w/v), temperature (30–50 °C), and particle size (250 μm–1.40 mm) affect hyper-ammonia-producing bacteria (HAB) growth during fermentation using commercially sourced pine wood-derived biochar. Fermentation experiments utilized enriched cow rumen fluid under controlled conditions, monitoring bacterial growth via optical density (OD600) over 48 h. Microbial proliferation was strongly influenced by all tested parameters (concentration, temperature, particle size). Highest growth occurred at 0.15% biochar concentration, 45 °C, and 250 μm particle size within the tested parameter ranges. Lower concentrations and smaller particles promoted microbial adhesion and colonization. Higher biochar levels hindered growth due to surface saturation and reduced pore accessibility. SEM imaging supported these findings by revealing structural changes on the biochar surface at different concentrations. Regression analysis demonstrated strong correlation between biochar parameters and microbial activity (R2 = 0.9931), though multicollinearity limited individual variable significance. These findings support biochar optimization for enhanced microbial processing in biotechnological applications.

1. Introduction

Microbial fermentation is influential across diverse industries, from producing foods and beverages to creating pharmaceuticals, biofuels, and biochemicals [1]. Fermentation processes rely on controlled microbial metabolism under anaerobic conditions to convert substrates into desired end products [2]. Although widely applied, microbial-induced reactions are inherently slow compared to chemical reactions [3]. Fermentation of complex feedstocks like lignocellulosic biomass, which resists breakdown without proper pretreatment, proceeds slowly [4]. Fermentation rates are constrained by microbial physiology and growth kinetics. Fermentative microbes are sensitive to medium compositions, time, temperature, and relative humidity, which can enhance or hinder their proliferation [5,6]. Creating a supportive environment for microbes can improve the fermentation process.
One approach is by immobilizing or attaching cells to a solid support matrix. Berillo et al. [7] explain that immobilizing microbial cells helps them maintain stability and concentration, which can lead to better yields and faster fermentation. This claim is supported by Najim et al. [8], who found that supported cultures generally offer better stress tolerance, reusability, and catalytic performance than free-floating cells. The enhanced adaptation of microbes in supportive environments has been demonstrated to improve fermentation efficiency by 20–40% compared to suspended cell systems [9]. However, choosing the right support materials remains a major challenge for industrial applications [10]. Synthetic resins, polymer gels, and activated carbon, while effective, can account for up to 30% of operational costs in large-scale fermentation systems [11]. To maintain economic viability and environmental sustainability, carrier materials must meet the criteria of affordability, availability, and renewability [12].
Researchers now explore natural, low-cost materials [13]. Biochar, produced by pyrolyzing biomass, is becoming a popular, affordable, and sustainable support material. By transforming agricultural residues into a valuable product, it promotes recycling and supports a circular economy. Additionally, biochar’s effects on microbial communities are generally positive, as studies have shown it does not harm microbes. Although microbes will react differently to biochar depending on the conditions, Benchaar et al. [14], the proliferation of hyper-ammonia-producing bacteria in the presence of biochar remains understudied.
Biochar’s porous structure, large surface area, and nutrient-rich particles offer a favorable environment for microbes to attach and thrive. It also contains functional groups and minerals like carbon, hydrogen, oxygen, and nitrogen, as well as key ratios like H/C and O/C [15,16]. The negatively charged surface of biochar helps attract nutrient ions and beneficial microbes [17]. Because biochar can create small, protective spaces that help microbes resist stress, it has proven to be an effective carrier for bacterial strains [18]. These traits make it an ideal environment for fermentative microbes, acting like a supportive scaffold that keeps them clustered and active. Biochar, as an affordable, environmentally friendly material, can be produced in large quantities and offers a porous structure well-suited for microbial growth [13].
Amending anaerobic microbial consortia with biochar led to high levels of biohydrogen yields from organic waste by encouraging biofilm formation and enriching beneficial bacteria [19]. Similarly, using biochar in ethanolic fermentation allowed microbes to handle normally inhibitive by-products, leading to significantly higher ethanol production under stress [20].
Hyper-ammonia-producing bacteria (HAB) represent a specialized group of microorganisms capable of producing elevated ammonia levels through enhanced amino acid deamination and protein degradation pathways [21]. Unlike conventional fermentative bacteria that primarily generate organic acids and alcohols, HAB maintain viability and drive ammonia production under elevated pH conditions (pH 7–11) [22]. Inclusion of biochar in the bio-ammonia fermentation system would enhance biofertilizer mineral composition and potentially serve as support for HAB proliferation. However, limited information exists about how biochar could affect HAB proliferation during fermentation, making this research important to pursue.
This study aims to explore how biochar properties and process parameters affect the growth of hyper-ammonia-producing bacteria (HAB) during fermentation, with the goal of identifying conditions that promote microbial proliferation and using SEM imaging to visually explore how biochar structure affects bacterial attachment and colonization.

2. Materials and Methods

The experimental workflow, from biochar preparation to fermentation setup and analysis is illustrated in Figure 1.

2.1. Media and Fermentation Conditions

The fistulated dairy cows used for rumen fluid collection are maintained at the North Dakota State University (NDSU) Animal Nutrition and Physiology Center. Rumen-derived hyper-ammonia-producing bacteria (HAB) were chosen for this study based on their demonstrated ability to efficiently degrade protein substrates while releasing ammonia with minimal energy input [23].
All animal procedures and rumen fluid collections were conducted in accordance with the ethical standards approved by the North Dakota State University Institutional Animal Care and Use Committee (IACUC) under protocol number IACUC20240021. The Animal Science Department, as the authorized handlers of the animals, oversaw all aspects of sample collection to ensure compliance with institutional and federal animal welfare guidelines.
The biochar used in this study was a commercial pine wood-derived product (Harris Horticultural Charcoal, Harris Products, LLC, Cartersville, GA, USA). The product is classified as premium raw biochar and is not pre-treated. The different particle size fractions were obtained through systematic mechanical sieving using ASTM standard sieves (ASTM E11 specifications) with vibratory separation for 30 min per fraction. Sequential sieving separated materials into defined fractions: <250 μm (collected on 250 μm sieve), 250–425 μm (retained between 425 μm and 250 μm sieves), and 425 μm–1.40 mm (retained between 1.40 mm and 425 μm sieves).
Fresh rumen fluid was collected from fistulated dairy cows and used within 24–48 h to minimize changes in microbial community composition and metabolic activity. Samples were stored at 39 °C (physiological temperature) in sealed 50 mL centrifuge tubes under anaerobic conditions to preserve the native microbial environment.
To cultivate and isolate hyper-ammonia-producing bacteria (HAB), an enriched medium was prepared following the formulation described by Bello et al. [22]. The medium per liter consisted of: Na2SO4 (0.480 g), NaCl (0.10 g), CaCl2·2H2O (0.064 g), Na2CO3 (4.0 g), K2HPO4 (0.292 g), KH2PO4 (0.292 g), yeast extract (0.50 g), and casamino acids (15 g). The pH was adjusted to 6.5 using HCl, and the medium was sterilized by autoclaving at 121 °C.
Prior to inoculation, the rumen fluid was allowed to settle, and the supernatant was diluted 1:100 to achieve an optical density (OD) of approximately 0.07. This diluted fluid was then added to the sterile medium and incubated at 39 °C with shaking at 130 rpm for 12 h. The microbial structures in the mimicry cultures were attributed to hyper-ammonia-producing bacteria (HAB) following the isolation protocol of Adeniyi et al. [23], which identifies Klebsiella quasivariicola, Escherichia coli, and Enterobacter cloacae as dominant HAB members in rumen-derived ammonia synthesis systems. The resulting culture was subsequently used as the inoculum for all experiments.

2.2. Experimental Setup

The fermentation experiments were conducted using a Thermo Scientific MaxQTM 4000 shaker (Thermo Fisher Scientific, Waltham, MA, USA), which ensured consistent agitation and precise temperature control. Each treatment was prepared in 250 mL sterile Erlenmeyer flasks containing 40 mL of autoclaved growth medium, inoculated with 0.4 mL of prepared HAB culture. Sterilized biochar was added according to the specific variable under investigation: concentration, temperature, or particle size. The flasks were securely clamped onto the shaker platform and incubated at 30–50 °C with a shaking speed of 130 rpm for 48 h, a timeframe commonly used in in vitro rumen studies to capture microbial activity without complications from nutrient depletion or metabolite accumulation [24]. These conditions promoted uniform mixing, prevented particle settling, and enhanced interaction between the biochar and microbes.
Samples were collected every six hours for optical density measurements at 600 nm (OD600) and filtered using Bacterial cells were separated from biochar particles using Whatman Grade 1 qualitative filter paper (11 μm retention) (Whatman International Ltd., Maidstone, UK). This filtration method ensures accurate optical density measurements by removing light-scattering biochar particles while preserving the bacterial suspension for analysis.

2.3. Biochar Concentration Experiments

The effect of varying biochar concentrations on the growth rates of HAB was evaluated using six different biochar loadings: 0.05%, 0.1%, 0.15%, 0.2%, and 0.25% (w/v). This range aligns with previous findings suggesting that low to moderate biochar levels can enhance microbial activity, while higher concentrations such as those above 2% reported by O’Reilly et al. [25], may reduce digestibility and hinder microbial function. Optical density (OD600) was measured at 6 h intervals over a 48 h incubation period using a spectrophotometer set at 600 nm. The growth medium was inoculated with HAB and incubated at 39 °C on a shaker set to 120 rpm.
A biochar particle size of 850 μm was selected to balance microbial accessibility, as supported by He et al. [26], who found that particles <1 mm enhance microbial colonization and sustained methanogenesis. To maintain consistency across treatments, 0.1 g of biochar was added to 40 mL of media, ensuring a representative biochar-to-liquid ratio, while 0.4 mL of HAB inoculum was used to provide a uniform microbial load for tracking growth patterns. All treatments were prepared in duplicates for reliability.
Bacteria growth phases were identified based on OD trends. Samples were collected and filtered through Whatman Grade 1 qualitative filter paper (11 μm retention), and the most favorable biochar concentration for bacterial growth was determined from the OD measurements.

2.4. Temperature Experiments

Experiments were conducted to evaluate the effect of temperature on the growth of HAB in the presence of biochar at five distinct temperatures: 30 °C, 35 °C, 40 °C, 45 °C, and 50 °C to systematically evaluate microbial adaptability, stability, and performance under non-rumen conditions. The study included two treatment groups: the first comprised medium and microbes only (MM), while the second comprised biochar, medium, and microbes (BMM). The MM treatments served as essential baseline controls to distinguish microbial growth in the absence of biochar from growth influenced by biochar addition. Each treatment was prepared in duplicate using 2 g of 850 μm biochar per 800 mL of media. For each experimental unit, 40 mL of media containing 0.1 g of biochar and 0.4 mL of HAB inoculum (for BMM only) was used. Samples were collected every 6 h for 48 h. OD was measured at each time point, and all samples were filtered using Whatman Grade 1 qualitative filter paper (11 μm retention) to remove biochar particles. A two-way ANOVA was conducted to assess the significance of differences in bacterial growth across treatments and temperatures.

2.5. Biochar Particle Size Experiments

The impact of biochar particle size on microbial interactions was evaluated using three biochar particle sizes: 250 μm, 425 μm, and 1.40 mm. Fine particles (<250 µm), especially when forming colloidal suspensions, pose notable challenges in bioprocessing: they are difficult to remove by standard filtration due to their stability and small size, which enables them to evade settling and pass through filters [27] and they contribute to fouling, cake layer formation, and flux decline [28].
Experiments were conducted in duplicate using growth media inoculated with HAB. Each treatment consisted of 40 mL of media, 0.1 g of biochar of the specified particle size, and 0.4 mL of HAB inoculum. Samples were collected every six hours over a 48 h period and filtered using Whatman Grade 1 qualitative filter paper (11 μm retention) to remove biochar particles. Microbial interaction levels were quantified by measuring OD values, and statistical analyses were performed to determine significant differences among particle sizes, thereby identifying the size that best supports microbial growth and activity.

2.6. Statistical Analysis

Two-way ANOVA was used to assess the effects of time and experimental variables, specifically biochar concentration, temperature, and particle size, on microbial growth. Statistical significance was determined at a p-value threshold of <0.05. All analyses were performed using R (version 4.3.0). Additionally, regression analysis was performed to evaluate the relationship between each experimental factor (biochar concentration, temperature, and particle size) and microbial growth, as measured by optical density (OD). This approach enabled us to identify conditions that are favorable for HAB proliferation within the tested parameter ranges.

2.7. Scanning Electron Microscope Analysis (SEM)

To prepare the samples, biochar was applied to adhesive carbon tabs that were subsequently mounted on cylindrical aluminum holders. To ensure optimal imaging conditions, the excess biochar was carefully removed using a nitrogen gas stream. Before imaging, a conductive gold coating was applied to the samples using a Cressington 108auto sputter coater (Ted Pella Inc., Redding, CA, USA) operating at 20 mA for 60 s. Images were obtained using a JEOL JSM-6490LV scanning electron microscope (JEOL USA, Peabody, MA, USA) at an accelerating voltage of 15 kV.

3. Results

3.1. Biochar Concentration Effects

The effect of biochar loading on HAB growth was evaluated over 48 h at 0.05–0.25% (w/v) (Figure 2). During the initial 6–18 h, OD600 increased across all treatments with no consistent separation by concentration. Divergence became evident by 24–30 h, with the 0.15% treatment exhibiting the highest OD600 in that interval. At 48 h, the 0.15% condition maintained a modest advantage, whereas higher loadings (0.20–0.25%) showed lower OD600 than the 0.05–0.15% range.
Across the concentration series, the abundance of free cells in suspension decreased as biochar loading increased, indicating a shift toward surface-associated biomass at higher loadings. Overall, 0.15% (w/v) yielded the strongest planktonic growth profile within the tested range.

3.2. Temperature Effects

HAB growth was evaluated across a temperature range of 30–50 °C over 48 h under two conditions: medium + HAB (MH) and biochar + medium + HAB (BMH) (Figure 3). At most temperatures, OD600 values were higher in BMH than in MH. The highest growth was recorded in the BMH treatment at 45 °C, with the maximum OD600 observed at 48 h.
At the lower end of the range, growth was reduced, with OD600 values of 0.567 at 30 °C and 0.572 at 35 °C at 48 h. Temperatures above 45 °C resulted in a decline in OD600 relative to the 45 °C condition.

3.3. Particle Size Effects

The effect of biochar particle size on HAB growth was assessed using 250 μm, 425 μm, and 1.40 mm fractions over a 48 h incubation (Figure 4). During the initial period (6–24 h), OD600 values were low and showed no clear separation among the treatments. By 30–48 h, differences became evident. The 250 μm and 425 μm treatments exhibited higher OD600 values relative to the 1.40 mm treatment, which consistently showed the lowest growth across all times.

3.4. Statistical Analysis Results

Two-way ANOVA indicated a strong effect of time on OD600 (F(1,56) = 782.970, p < 2 × 10−16) and a significant effect of experimental condition (biochar concentration, temperature, particle size) (F(11,56) = 8.427, p = 1.65 × 10−8). A significant time × condition interaction was also observed (F(11,56) = 3.958, p = 0.000285), showing that growth trajectories varied across treatments. Regression analysis incorporating concentration, temperature, particle size, and their interactions explained 99.31% of the variance in OD600 (R2 = 0.9931; adjusted R2 = 0.9758). None of the individual predictors or interaction terms were statistically significant (p > 0.05).

3.5. Biochar, Microbe, and Composite Morphology

SEM imaging was performed to examine biochar morphology and microbial attachment at different concentrations (0.05–0.25% w/v) (Figure 5). Pristine pine-derived biochar displayed a porous structure with open channels and well-defined cavities (Figure 5a). At 0.05–0.15% (Figure 5b–d), microbial cells were visible along pore walls and within surface cavities, indicating colonization at these loadings.
At 0.20–0.25% (Figure 5e,f), the biochar surface appeared more compact, with evidence of pore occlusion and particle aggregation. Microbial cells were still observed, but attachment appeared less uniformly distributed compared with the lower concentrations.

4. Discussion

4.1. Scientific Novelty and Mechanistic Basis

This study is the first systematic evaluation of how biochar concentration, particle size, and temperature jointly influence the proliferation of hyper-ammonia-producing bacteria (HAB). The results show that immobilization outcomes are determined by the combined influence of matrix properties and process conditions rather than by single parameters.
Consistent with previous reports that no universal biochar concentration exists [29], HAB growth peaked at a moderate loading (0.15% w/v), while higher concentrations (0.20–0.25%) inhibited proliferation. Similar inhibitory effects have been reported, where excessive biochar suppressed microbial activity through substrate saturation and nutrient immobilization [30,31,32]. SEM analysis confirmed that biochar provided a porous and structurally favorable substrate for colonization [33], supporting the well-documented role of biochar in microbial attachment [34,35].
The study also clarifies the impact of particle size, a parameter often overlooked in biochar–microbe interactions. Smaller particles are known to enhance colonization through greater surface area [36,37,38,39], with positive effects on fermentation efficiency and digestibility [40,41]. Wu et al. [42] reported that fine particles facilitate microbial interactions by improving electron transfer, while larger particles are less effective due to limited accessibility [43]. In agreement with these findings, HAB colonization in this study was strongest at 250–425 μm, whereas the 1.40 mm fraction consistently supported lower growth.
Temperature was identified as a second major determinant of HAB performance. Microbial growth is strongly influenced by thermal conditions [33,34], with elevated levels accelerating turnover in some systems [44] but imposing stress in others [35,36,45]. HAB maintained growth between 30 and 50 °C, with maximum proliferation at 45 °C. This adaptability is favorable for application under variable environmental conditions and is consistent with evidence that biochar stabilizes enzymes and supports microbial activity under heat stress [37,38,39].
These findings indicate that HAB physiology optimized for ammonia production and alkaline pH tolerance [22,23] requires biochar designs distinct from DIET- or hydrolysis-oriented systems developed for methanogens and acidogens [42,46]. The integration of growth kinetics, SEM evidence, and statistical analysis broadens the scope of biochar–microbe studies to include ammonia-producing consortia, a microbial group that has received limited attention.

4.2. Practical Implications and Environmental Considerations

The combination of HAB with biochar has important implications for nitrogen management. Biochar-based systems are known to reduce nitrogen losses through lower volatilization and leaching [15,16,37,39], and field studies have reported 25–40% reductions compared with conventional fertilizers [37,39]. Incorporating HAB into such systems adds a biological dimension, providing ammonia production synchronized with microbial metabolism, which complements the physical retention capacity of biochar.
This dual functionality biological nitrogen provision and carbon sequestration aligns directly with sustainable agriculture priorities. From an economic perspective, HAB–biochar systems may be particularly advantageous for organic production, where higher premiums on inputs can offset processing costs [47,48,49]. In addition, biologically mediated nitrogen release presents a lower environmental risk than concentrated synthetic fertilizers, as microbial activity adjusts to soil conditions and plant demand [50,51]. Biochar further contributes to environmental benefits by reducing nitrate leaching and nitrous oxide emissions, thereby reinforcing the sustainability potential of the HAB–biochar approach.

4.3. Limitations and Future Research Directions

Although this study establishes clear links between biochar properties and HAB proliferation, several limitations remain. The parameter ranges investigated were relatively narrow, and more comprehensive experimental designs, such as response surface methodology or central composite designs, will be required to define optimal conditions more precisely. Additional characterization of the biochar, including pore size distribution, hydrophobicity, and surface chemistry, is also needed to quantitatively relate physical structure to microbial attachment [33,44].
The experiments were restricted to short-term growth assays. Longer-term studies are necessary to evaluate biofilm formation, persistence, and colonization stability under repeated or extended use. Moreover, field trials will be essential to validate laboratory findings under agricultural conditions, particularly across seasonal variation in soil temperature, moisture, and crop demand. Finally, integration of HAB inoculation into existing biochar production systems will be critical for assessing scalability and commercial feasibility for biofertilizer development.

5. Conclusions

This study systematically evaluated how biochar parameters (concentration and particle size) and a process condition (temperature) influence the growth of hyper-ammonia-producing bacteria (HAB). The results showed that HAB proliferation was greatest at a biochar concentration of 0.15% (w/v), was enhanced with smaller particle sizes (250–425 μm), and reached its optimum at a temperature of 45 °C. SEM imaging confirmed that biochar surfaces supported microbial colonization across treatments, while higher concentrations reduced the abundance of free cells in suspension.
The novelty of this work lies in demonstrating that HAB, unlike conventional fermentative bacteria, require specific combinations of concentration, particle size, and temperature to achieve optimal immobilization and growth. These findings establish a framework for tailoring biochar-based systems to the unique physiology of HAB and provide a foundation for the development of biofertilizer platforms based on HAB–biochar composites.
Future research should extend these findings by applying advanced experimental designs, incorporating detailed biochar characterization, and validating the system under field conditions to assess persistence, scalability, and agricultural performance.

Author Contributions

Conceptualization, A.H., C.B. and E.M.; methodology, C.B., A.H. and T.A.; validation, A.H.; formal analysis, C.B., A.H. and T.A.; investigation, C.B.; resources, A.H.; data curation, C.B. and T.A.; writing—original draft preparation, C.B.; writing—review and editing, A.H., T.A., K.A., N.C.S. and C.C.; visualization, C.B. and T.A.; supervision, A.H.; project administration, A.H.; funding acquisition, A.H. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by North Dakota Soybean council, ND, U.S.; Grant number FAR0038184.

Institutional Review Board Statement

The animal study protocol was approved by the Institutional Review Board (or Ethics Committee) of the North Dakota State University Institutional Animal Care and Use Committee (IACUC) (protocol code IACUC20240021 and date of approval: 17 April 2024).

Data Availability Statement

The datasets generated and analyzed during the current study are available from the corresponding author upon reasonable request. All processed data supporting the findings are included within this article.

Acknowledgments

The authors acknowledge the North Dakota State University Animal Nutrition and Physiology Center for providing access to fistulated dairy cows and technical support. We thank the USDA-ARS NCSL NDSU Electron Microscopy Core for assistance with SEM imaging.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. Process Flowchart for Biochar-Based Microbial Fermentation Study.
Figure 1. Process Flowchart for Biochar-Based Microbial Fermentation Study.
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Figure 2. Growth trends of HAB over 48 h at varying biochar concentrations (0.05–0.25% w/v).
Figure 2. Growth trends of HAB over 48 h at varying biochar concentrations (0.05–0.25% w/v).
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Figure 3. Effect of temperature (30–50 °C) on HAB growth in the presence of biochar.
Figure 3. Effect of temperature (30–50 °C) on HAB growth in the presence of biochar.
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Figure 4. Effect of biochar particle size (250 μm, 425 μm, and 1.40 mm) on HAB growth over 48 h.
Figure 4. Effect of biochar particle size (250 μm, 425 μm, and 1.40 mm) on HAB growth over 48 h.
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Figure 5. SEM micrographs of pine wood-derived biochar at different concentrations showing HAB colonization. (a) Pristine biochar; (b) 0.05% w/v; (c) 0.10% w/v; (d) 0.15% w/v; (e) 0.20% w/v; (f) 0.25% w/v. Red arrows indicate attached HAB cells, while circles depict pore clogging and reduced microbial visibility.
Figure 5. SEM micrographs of pine wood-derived biochar at different concentrations showing HAB colonization. (a) Pristine biochar; (b) 0.05% w/v; (c) 0.10% w/v; (d) 0.15% w/v; (e) 0.20% w/v; (f) 0.25% w/v. Red arrows indicate attached HAB cells, while circles depict pore clogging and reduced microbial visibility.
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MDPI and ACS Style

Bitrus, C.; Hammed, A.; Ayodele, T.; Alarape, K.; Chandra Sarker, N.; Clementson, C.; Monono, E. Adopting Biochar as Immobilization Support for Hyper Ammonia-Producing Bacteria Proliferation. Appl. Microbiol. 2025, 5, 111. https://doi.org/10.3390/applmicrobiol5040111

AMA Style

Bitrus C, Hammed A, Ayodele T, Alarape K, Chandra Sarker N, Clementson C, Monono E. Adopting Biochar as Immobilization Support for Hyper Ammonia-Producing Bacteria Proliferation. Applied Microbiology. 2025; 5(4):111. https://doi.org/10.3390/applmicrobiol5040111

Chicago/Turabian Style

Bitrus, Christiana, Ademola Hammed, Tawakalt Ayodele, Kudirat Alarape, Niloy Chandra Sarker, Clairmont Clementson, and Ewumbua Monono. 2025. "Adopting Biochar as Immobilization Support for Hyper Ammonia-Producing Bacteria Proliferation" Applied Microbiology 5, no. 4: 111. https://doi.org/10.3390/applmicrobiol5040111

APA Style

Bitrus, C., Hammed, A., Ayodele, T., Alarape, K., Chandra Sarker, N., Clementson, C., & Monono, E. (2025). Adopting Biochar as Immobilization Support for Hyper Ammonia-Producing Bacteria Proliferation. Applied Microbiology, 5(4), 111. https://doi.org/10.3390/applmicrobiol5040111

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