Post-Inoculation Drying and Storage Effects on HAB Viability and Nutrient Retention in Biochar

TL;DR

This study explores how drying temperatures affect the survival and recovery of beneficial bacteria in biochar and their impact on nutrient retention.

Contribution

The study reveals that higher drying temperatures initially reduce microbial viability but may enhance recovery after storage.

Findings

Higher drying temperatures reduced immediate microbial revival but improved recovery after 30 days of storage.

Drying temperature altered surface functional groups linked to microbial attachment and activity.

Nitrogen retention in biochar was minimally affected by drying temperatures above 55 °C.

Abstract

Background/Objectives: The effects of thermal drying on the viability of beneficial microorganisms immobilized in biochar, as well as on biochar nutrient retention, remain insufficiently understood. This study aimed to evaluate how drying temperature influences the survival of hyper-ammonia-producing bacteria (HAB) immobilized on pine wood biochar and to assess the impact of subsequent storage on bacterial recovery and nutrient stability. Methods: Biochar was inoculated with HAB and subjected to drying at temperatures ranging from 40 to 60 °C. Following drying, samples were characterized and stored for 30 days. Microbial revival was assessed through reculturing, while changes in surface functional groups were analyzed using FTIR spectroscopy. Nutrient retention, particularly nitrogen content, was also evaluated. Results: Higher drying temperatures resulted in reduced immediate microbial revival during reculturing. However, samples exhibiting limited immediate recovery demonstrated enhanced revival after the 30-day storage period. FTIR analysis revealed that drying temperature modified the availability of surface functional groups associated with microbial attachment and activity. Nutrient analysis indicated only minor reductions in nitrogen retention in biochar dried at temperatures above 55 °C. Conclusions: Drying temperature significantly affects both the short-term survival and post-storage recovery of beneficial microorganisms immobilized in biochar. While elevated temperatures may initially suppress microbial activity, recovery potential during storage remains substantial. Optimizing drying conditions is therefore essential to balance microbial viability with nutrient retention in biochar-based formulations.

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Full text

1. Introduction

Nitrogen (N) management has been one of the most enduring problems in current agricultural practices due to inefficient fertilizer use, causing both environmental damage and financial loss. Conventional synthetic N fertilizers exhibit low nitrogen use efficiency, and an estimated 50–70% of applied N is lost through volatilization, leaching, and runoff [1,2,3,4]. These losses result in increased ammonia emission into the atmosphere, increased nitrates contaminating groundwater, increased eutrophication and greenhouse gas production [5,6,7].

The potential benefits of losing less N may be worth over $400 billion per year, and as stated in [8], there is a significant need for better methods of delivering N efficiently and sustainably. Given these limitations of conventional nitrogen fertilization, attention has increasingly turned to soil amendments capable of enhancing nutrient retention and cycling while mitigating associated environmental impacts.

Biochar is known to be one of the most promising amendments used to improve soil fertility, thanks to its stable properties, porous structure and wide variety of chemical functional groups [9,10,11]. The recalcitrant carbon matrix in biochar provides an improved cation exchange capacity, better soil structure, better moisture retention, and a protective environment for microbes [12,13,14,15]. Based on meta-analysis studies, adding biochar increases microbial biomass and microbial activity; however, the effect is dependent upon the type of biochar added and the characteristics of the soil [16,17,18].

The combination of biochar with plant growth-promoting microorganisms (PGPMs), nitrogen-fixing bacteria, or other nutrient-solubilizing microorganisms may also increase the efficiency of nitrogen cycling processes by creating a favorable environment for microbial activity. The incorporation of biochar into soils creates sites where ammonia can be adsorbed, thus reducing the volatilization of ammonia and improving the retention of ammonium. This process has been shown to produce both enhanced microbial populations and increased levels of nutrients available to plants [19,20,21]. In spite of this rapid expansion of the biofertilizer industry, there are still a number of commercial microbial inoculants that have varying degrees of success in the field due to issues associated with the drying out of microorganisms, temperature changes, and degradation of the carrier material used during storage and transportation [22,23]. Biochar’s unique structural and chemical characteristics provide an opportunity to create more stable environments for microorganisms; however, relatively little research has investigated the effect of stabilizing the physical/chemical properties of biochar on the viability of microorganisms after the inoculum has been introduced into the soil, particularly for fast-ammonifying organisms like HAB.

HAB have historically been studied in ruminant microbiology [24,25] and are known to have a very high efficiency in their deamination pathways, which create significant amounts of ammonium when grown under anaerobic conditions [26]. Recent studies suggest that HAB may be used as microbial ammonium-producing microorganisms if they can be immobilized within a solid matrix [27]. Therefore, the incorporation of HAB into a solid matrix such as biochar could allow for both the biological production of ammonium through ammonification and a mechanism for retaining NH_4_^+^ in the soil, providing a biologically active slow-release source of nitrogen fertilizer.

One of the challenges of creating a microbial fertilizer using biochar lies in the post-inoculation process of stabilizing the microbial culture. This includes drying, curing and storing the cultures. The drying process can cause damage to the bacterial cell membrane, reduce the culturable population of bacteria and affect how the bacteria metabolize [28,29]; at the same time, the curing process causes physical and chemical changes to occur in the biochar. For example, there are losses of the surface functional groups in the biochar, collapses of the pores within the biochar, and pH changes, which may result in less sorption of nutrients and create different microbial habitats [30,31]. Survival of microbes in biochar varies widely based on many factors, including the type of feedstock used, the structure of the pores, the types of microbes, and the environmental conditions during storage [32,33,34]. Loss of moisture from the biochar causes the bacteria to go dormant or encourages the growth of bacteria that are tolerant to water loss, and this could lead to inconsistencies in the performance of the product in the field [35,36,37,38,39].

Despite extensive research on the agronomic value of biochar and the plant growth benefits of hydrolytic and associative bacteria (HAB), very limited attention has been given to the post-inoculation stabilization phase that occurs immediately after microbial enrichment. Existing studies focus primarily on biochar production temperature, feedstock type, or microbial inoculation strategies but rarely examine how drying conditions applied after inoculation influence microbial survival, nutrient retention, or functional group integrity. Moreover, while biochar is widely promoted as a carrier for microbial inoculants, the optimal temperature range that preserves both microbial viability and biochar functionality during drying and storage remains largely unresolved. This represents a critical knowledge gap, as inappropriate post-inoculation stabilization may severely reduce the shelf life and field effectiveness of biochar-based biofertilizers. The aim of this study was to determine how post-inoculation drying temperature and subsequent storage affect the viability and recovery of hyperammonia-producing bacteria (HAB) immobilized on biochar over time while simultaneously assessing changes in functional groups and nutrient content. By integrating microbial survival kinetics with FTIR and nutrient analyses, this work provides the first systematic assessment of post-inoculation stabilization conditions for HAB-enriched biochar, offering practical guidance for developing more robust and storage-stable microbial biofertilizer formulations.

2. Materials and Methods

2.1. Media and Fermentation Conditions

Modifications of fermentation procedures used in [27] were made to address the objectives of this research study. A nutrient-rich basal medium was prepared to support nitrogen-transforming microbes through their metabolic processes. The basal medium was sterilized using an autoclave at 121 ° C for 15 min to create a sterile environment. The inoculum used for fermentation was an active culture of a previously isolated, anaerobically grown, hyperammonia-producing bacterium (HAB). These bacteria were chosen because of their potential to produce ammonia and as a candidate for biofertilizer applications. The fermentation process was conducted at optimal temperatures and pH levels for HAB growth and the release of ammonium ions.

2.2. Fermentation Procedure

The Thermo Scientific MaxQ™ 6000 orbital shaker (Thermo Fisher Scientific, Waltham, MA, USA) was selected for use in fermentation studies due to it being able to maintain both stable temperatures and a uniform motion throughout the duration of the study. Each experiment was conducted in a 100 mL sterile GL 45 glass media bottle (VWR International, Radnor, PA, USA), containing 40 mL of autoclaved media that had been enriched with nutrients. Initiation of fermentation occurred through the aseptic transfer of 0.4 mL of an actively growing HAB culture into each flask, followed by the addition of 2 g of sterilized pine wood-based biochar. The flasks were then securely fastened onto the orbital shaker and maintained under constant agitation at 130 rpm and at a temperature of 45 °C for a period of 12 h. This operational configuration was established to ensure consistent blending of all components, to minimize settling of the biochar, and to optimize the interaction of the bacterial surface with the biochar.

Samples were obtained at 12 h intervals to determine microbial growth based upon the optical density of the solution at a wavelength of 600 nm (OD_600_) using a Tecan Infinite^®^ Nano+ spectrophotometer (Model 200 Pro, Tecan Austria GmbH, Grödig, Austria). Bacterial cells contained within the biological sample were removed from contact with the biochar particles prior to spectrophotometric measurement through filtration of the entire sample using Whatman Grade 1 Qualitative Filter Paper (11 µm retention; Whatman International Ltd., Maidstone, UK). The biochar particles were retained in the filter and therefore eliminated as potential light scatterers during the spectrophotometric measurements of the bacterial cells. All inoculation steps were carried out under a laminar-flow hood using autoclaved glassware and sterile instruments to prevent contamination. Media blanks were incubated alongside experimental flasks to confirm sterility throughout the fermentation process.

2.3. Replication and Experimental Design

All microbial viability and OD_600_ measurements were performed using two independent biological replicates per temperature treatment (n = 2). Each biological replicate consisted of a separately inoculated biochar sample that underwent the assigned drying treatment and was stored under identical conditions. For OD_600_ measurements, two technical replicate readings were taken for each biological replicate and averaged. All results are reported as means ± standard deviation (SD).

2.4. Post-Fermentation Processing

The filtered liquid was removed from the biochar/liquid mixture with Whatman Qualitative Filter Paper to allow for the recovery of the biochar portion. Following this filtering step, the recovered biochar fractions were dried (using an oven) for 24 h, at temperatures ranging between 40 °C and 60 °C; these temperatures are representative of typical post-inoculum stabilization regimes that are typically used in preparation of biofertilizers for formulation, storage, etc., thereby allowing reductions in moisture content while maintaining the viability of microorganisms present on the surface of the biochar. After drying, the biochar fractions were placed into airtight, sterile containers and held at room temperature until further physicochemical and microbiological analysis could be conducted.

2.5. Assessment of Microbial Viability After Drying

To evaluate the recovery of immobilized microorganisms after the drying phase, each dried biochar sample (0.1 g) was aseptically transferred into fresh medium and incubated under controlled conditions for 48 h. Microbial growth during reculturing was monitored by measuring OD_600_ at 12 h intervals using a Tecan Infinite^®^ Nano+ spectrophotometer (Model 200 Pro, Tecan Austria GmbH, Grödig, Austria). This assay assessed the ability of immobilized HAB to revive and regain metabolic activity following drying. OD_600_ measurements in the reculturing assay were used as indicators of functional revival capacity rather than direct quantification of viable cell numbers. An increase in OD_600_ during reculturing requires intact cellular membranes, preserved metabolic machinery, and the ability to resume growth under favorable conditions. Therefore, the recovery dynamics reported in this study reflect the capacity of immobilized HAB populations to regain metabolic activity following drying and storage, rather than absolute viable counts.

Colony-forming unit (CFU) enumeration was not feasible because HAB cells strongly adhered to and embedded within the biochar pore matrix. Detachment methods such as vortexing, sonication, or chemical dispersants may selectively recover loosely attached cells while underestimating protected subpopulations or introducing membrane damage. Consequently, CFU counts could introduce greater methodological bias than reculturing-based revival assays in this carrier system.

2.6. Fourier Transform Infrared Spectroscopy (FTIR)

FTIR analysis was performed to identify changes in surface functional groups following HAB inoculation and temperature-dependent stabilization. Approximately 1 g of finely ground biochar was mixed with spectroscopic-grade KBr at a 1:100 (w/w) ratio and pressed into pellets. Spectra were collected using a Nicolet 8700 FTIR spectrometer (Thermo Fisher Scientific, Madison, WI, USA) operated with OMNIC™ software (OMNIC SST 8.1, Thermo Fisher Scientific) over 4000–400 cm^−1^ at 4 cm^−1^ resolution, averaging 32 scans per sample. An empty-beam (air) background spectrum was collected prior to sample acquisition and automatically subtracted by the software. Spectra were baseline-corrected prior to interpretation.

A separate pure KBr pellet spectrum was not collected during these measurements. However, spectroscopic-grade KBr is largely transparent in the mid-infrared region and does not exhibit characteristic absorption bands within the functional group regions evaluated in this study (3000–3500, ~1700, and 1000–1300 cm^−1^). Therefore, the absorption bands reported are attributable to the biochar samples rather than the KBr matrix. Biochar functional group assignments were based on comparison of peak positions with previously published reference spectra for biochar materials. For quantitative analysis, spectra originally collected in transmittance mode were exported and converted to absorbance using the relationship A = log_10_(1/T), where T represents transmittance. Integrated peak areas were calculated over fixed wavenumber intervals corresponding to O–H stretching (3000–3500 cm^−1^), C–H stretching (2850–2960 cm^−1^), C=O stretching (~1650–1750 cm^−1^), and C–O/C–OH vibrations (1000–1300 cm^−1^). Identical integration limits were applied to all samples to ensure direct comparability across treatments.

2.7. Total NPK

Total nutrient content of the biochar samples, including total nitrogen (N), phosphorus (reported as phosphate, P_2_O_5_), and potassium (reported as potash, K_2_O), was analyzed by Agvise Laboratories (Northwood, ND, USA) using standard soil and fertilizer analytical procedures. Total N was determined by combustion (Dumas method). Phosphorus and potassium were determined following acid digestion/extraction and quantified by inductively coupled plasma (ICP) analysis. All nutrient values represent means ± SD of duplicate laboratory analyses and are reported on an as-received basis.

2.8. Storability

After inoculating the biochar with HAB, we placed the biochar–HAB formulation in a stabilization environment where it was maintained for a period prior to storing it in a sterilized centrifuge tube at room temperature to mimic the conditions that are typical of biofertilizers when they are stored. The samples were then removed from the tubes after thirty days of storage to assess the viability and longevity of the hyperammonia-producing bacteria (HAB) in the biochar formulation. We used a reculture method on casamino acid-enriched medium to assess microbial survival, in which 0.1 g of the stored biochar was added to the culture and allowed to grow at optimal temperatures and conditions. After allowing the culture to incubate and grow, we filtered the culture to eliminate residual particulate matter associated with the biochar and measured microbial growth by spectrophotometry at 600 nm (OD_600_), providing an objective measure of HAB viability and metabolic activity after extended periods of storage that reflect the shelf-stable nature of the biochar formulation as a biofertilizer. No microbiological measurements were collected during the storage period; viability was evaluated only at the end of storage using the standardized recovery (OD_600_) assay.

3. Results

3.1. Drying at Different Temperatures

Drying temperature had a clear influence on the immediate revival of HAB following inoculation (Figure 1). At 12 h, OD_600_ values were low across all treatments, ranging from 0.006 at 50 °C to approximately 0.032 at 40–55 °C. By 24 h, the highest OD_600_ increase occurred at 50 °C (0.095), followed by 60 °C (0.047) and 55 °C (0.040), whereas 45 °C showed limited revival (~0.019). The strongest revival occurred at 36 h, where 50 °C and 40 °C exhibited the highest OD_600_ values (0.103 and 0.102, respectively). In contrast, 45 °C remained consistently low (~0.021), indicating temperature sensitivity. By 48 h, OD_600_ declined slightly at 50 °C (0.061) and 40 °C (0.063), while 55 °C showed continued growth (0.073). The 60 °C treatment exhibited minimal recovery throughout, decreasing from 0.047 at 24 h to only 0.007 at 48 h; 50 °C supported the strongest short-term viability, followed closely by 40 °C, while temperatures ≥ 55 °C suppressed revival, and 60 °C resulted in near-complete loss of immediate metabolic activity.

3.2. Storability of Inoculated Biochar

After 30 days of storage, HAB revival patterns differed markedly from the immediate post-drying response (Figure 2). At 12 h of reculturing, the highest OD_600_ value occurred at 55 °C (≈0.081), followed by 60 °C (≈0.052) and 50 °C (≈0.033), whereas 40 °C and 45 °C showed lower revival (≈0.027 and ≈0.006, respectively). By 24 h, modest increases were observed at 40 °C (≈0.038), 55 °C (≈0.035), and 60 °C (≈0.031), while 45 °C (≈0.017) and especially 50 °C (≈0.004) remained low. Thus, early-stage recovery after storage was strongest at 40–60 °C, with 45–50 °C showing the poorest growth.

A strong divergence appeared at 36 h, when 45 °C and 60 °C exhibited the highest OD_600_ values (≈0.092 and ≈0.091, respectively), clearly exceeding those at 40 °C and 55 °C (both ≈0.030) and 50 °C (≈0.011). This pattern indicates a delayed but pronounced revival potential in cells exposed to moderate or high drying temperatures. By 48 h, 60 °C reached the highest OD_600_ (≈0.128), followed by 55 °C (≈0.065) and 45 °C (≈0.033). In comparison, 40 °C showed limited final growth (≈0.011), and 50 °C remained consistently low (≈0.023), indicating poor recovery after storage. Storage fundamentally altered the viability ranking observed after drying; whereas 40–50 °C supported the strongest immediate revival, the highest post-storage recovery occurred at 55–60 °C, suggesting the presence of stress-tolerant or VBNC-like subpopulations that regained activity only after prolonged reculturing.

3.3. Nutrient Profiles Post-Inoculation

Nutrient concentrations of the HAB-inoculated pine wood biochar were evaluated across the 40–60 °C post-inoculation drying treatments and are reported on an as-received basis.

Across the 40–60 °C drying range, nutrient concentrations in HAB-inoculated pine biochar showed overall stability, with only minor temperature-associated variation (Table 1). Total nitrogen (N) was highest at 40 °C (1.37%) and decreased slightly at higher temperatures, reaching 1.19–1.21% at 50–60 °C. Although the decline is small, this pattern suggests that higher drying temperatures may contribute to modest nitrogen loss or reduced N retention, potentially due to volatilization of ammoniacal-N or reduced adsorption capacity as biochar dries more aggressively. However, the relatively narrow N range across treatments (mean ± SD = 1.24 ± 0.07%) indicates that N remained largely conserved even at 60 °C.

In contrast, phosphate (P_2_O_5_) showed minimal sensitivity to drying temperature, remaining within a narrow band (0.81–0.87%) across all treatments (mean ± SD = 0.84 ± 0.03%). This consistency indicates that phosphorus was highly stable during post-inoculation drying, likely because P is less prone to thermal losses under these conditions and may remain bound to biochar surfaces or retained in microbial-derived residues. Potash (K_2_O) exhibited the least variability, with nearly uniform values of 0.35% for 45–60 °C and only a small increase at 40 °C (0.40%). The weak temperature response suggests potassium retention was not strongly influenced by stabilization temperature within this range, consistent with the generally non-volatile nature of K under mild drying regimes. These results indicate that while drying temperature can slightly affect nitrogen retention, particularly above 50–55 °C, P and K concentrations remain essentially unchanged, supporting the conclusion that nutrient retention in HAB-enriched biochar is generally robust across commonly used post-inoculation drying temperatures.

3.4. FTIR Spectral Analysis

The FTIR spectra of the pine biochar enriched with HAB (Figure 3) show clear temperature effects on surface functional groups that are associated with nutrient binding and microbial attachment. All samples exhibited characteristic biochar peaks, including broad O–H stretching between 3200 and 3500 cm^−1^, aliphatic C–H stretching at 2850–2960 cm^−1^, C=O stretching around 1700 cm^−1^, and C–O/C–OH vibrations between 1000 and 1300 cm^−1^.

To provide quantitative support for the observed spectral trends, integrated peak areas were calculated following conversion of transmittance to absorbance. Integration of the O–H region (3000–3500 cm^−1^) showed a general decrease at higher drying temperatures, with the lowest values observed at 55 °C. In contrast, the 1000–1300 cm^−1^ region exhibited a gradual increase in integrated area from 17.7 at 40 °C to 20.2 at 60 °C. Integrated values are summarized in Table 2.

These quantitative results support the visual trends observed in the stacked spectra. At 40–45 °C, spectra showed the strongest O–H features, indicating preservation of hydroxyl functionalities. Drying at 50 °C resulted in a moderate reduction in O–H intensity, while more pronounced reductions were observed at 55 °C. The 60 °C treatment exhibited diminished O–H intensity relative to lower temperatures, consistent with thermal alteration of oxygen-containing surface groups.

At 55 °C, reductions were more pronounced in the O–H region, and the C=O peak at ~1700 cm^−1^ showed reduced intensity. The 60 °C treatment exhibited the greatest overall alteration in oxygen-containing functional groups. These temperature-dependent chemical changes help explain the observed differences in immediate microbial recovery while highlighting that functional group preservation alone does not fully determine long-term revival capacity.

4. Discussion

Drying temperature exerted a clear and substantial influence on the immediate culturability and subsequent revival of HAB immobilized on pine wood biochar, reflecting the intertwined effects of thermal stress, microbial physiology, and biochar microhabitat characteristics. Immediately after drying, the samples treated at 40–50 °C retained the highest culturability, whereas marked declines occurred at temperatures ≥ 55 °C. This pattern aligns with established cellular responses to heat stress, where moderate temperatures help maintain membrane fluidity and protein stability, while higher temperatures accelerate irreversible denaturation, oxidative damage, and loss of culturability [41,42]. The porous and chemically heterogeneous structure of biochar likely provided microsites that buffered HAB cells from complete desiccation injury, consistent with prior reports that biochar enhances microbial persistence under environmental stress [43,44].

Contrary to the immediate post-drying results, long-term recovery patterns after 30 days of storage shifted considerably. Although the 40–45 °C treatments continued to support moderate revival, the 50 °C samples, which had displayed the highest initial culturability, showed reduced recovery after storage. In contrast, the 55 °C and 60 °C treatments exhibited substantial OD_600_ increases during reculturing, despite showing minimal initial revival. These patterns are consistent with VBNC-like behavior or dormancy induced by heat and desiccation stress, in which cells temporarily lose culturability but remain capable of resuscitation under favorable conditions [45,46]. Similar recovery dynamics have been associated with stress-adaptation responses such as heat-shock protein induction, oxidative stress tolerance, and restoration of membrane integrity during rehydration and regrowth [28,29]. Moreover, porous carrier materials such as biochar can promote persistence of stressed subpopulations by reducing desiccation injury and providing protected microhabitats that support long-term revival [43,44]. However, because OD_600_ reflects recovery dynamics rather than direct cell counts, VBNC status cannot be confirmed without complementary viability-based assays. Because intermediate monitoring during storage was not performed, the timing of viability loss or recovery dynamics during storage cannot be resolved, and only the end-point recovery after 30 days is reported. This pattern may also reflect thermal conditioning, in which exposure to acute but sublethal heat induces protective mechanisms such as heat-shock protein expression, membrane phase adjustments, or DNA repair pathways. These adaptations enhance long-term endurance even if short-term culturability is compromised. Biochar’s structural diversity may further support the survival of these stress-tolerant subpopulations by offering hydrophobic microdomains, micropores, and surfaces that reduce water loss and slow oxidative damage. Because HAB cells strongly adhered to the biochar matrix, colony-forming unit (CFU) enumeration was not feasible; thus, OD-based revival assays were used to assess trends in functional revival capacity. Future studies should directly test whether HAB enters a VBNC or dormancy state under high-temperature drying by incorporating complementary viability-based analyses. These include membrane integrity assays (e.g., live/dead fluorescence staining), qPCR-based quantification to distinguish persistence from loss of viable cells, and evaluation of stress-response pathways such as heat-shock protein expression. These approaches would allow confirmation of whether delayed recovery reflects reversible loss of culturability rather than irreversible cell death.

Parallel changes in biochar surface chemistry help contextualize these microbiological outcomes. FTIR spectra showed progressive reductions in hydroxyl, carbonyl, and C–O/C–OH functional groups as drying temperature increased. Loss of these oxygen-containing groups is consistent with dehydration and partial oxidation of surface functionalities [47,48]. These groups play critical roles in moisture retention, microbial adhesion, nutrient sorption, and biofilm development. Their preservation at lower temperatures supports the stronger initial viability at 40–50 °C, whereas their degradation at ≥55 °C likely contributes to reduced immediate culturability. Nevertheless, the ability of HAB populations in the 55–60 °C treatments to recover after storage indicates that functional group degradation does not preclude microbial persistence, especially when VBNC-like responses or dormancy and biochar-mediated microhabitat protection are present. Changes in nutrient retention further support the observed interactions between microbial and chemical responses. Drying temperature had a modest effect on total N, which decreased slightly as temperature increased. This trend aligns with FTIR results showing reduced intensity of oxygen-containing functional groups (e.g., hydroxyl and carboxyl), suggesting fewer polar reactive sites for retaining nitrogen species introduced during inoculation. In contrast, phosphate (P_2_O_5_) and potash (K_2_O) remained stable across treatments, consistent with FTIR evidence that mineral-associated binding environments were largely unaffected within the 40–60 °C drying range.

Despite these shifts, all inoculated samples exhibited higher N, P, and K concentrations compared with uninoculated pine biochar [40], reflecting the nutrient contributions associated with the HAB inoculation process itself. Thus, even as surface chemistry shifted at higher temperatures, biochar retained substantial nutrient value.

The HAB–biochar formulations exhibited a multi-phase response to thermal stabilization, showing that immediate culturability alone does not reliably predict long-term survival. These findings show the importance of evaluating both immediate and post-storage microbial performance when designing and optimizing biochar-based microbial carriers. Although OD_600_ effectively captured the growth trends and revival kinetics of the HAB populations, it is acknowledged that absorbance readings can be influenced by background signals from biochar particulates. In stress tolerance and desiccation studies, recovery-based assays are commonly employed to evaluate the capacity of bacterial populations to resume growth following environmental perturbation [28,29,45,46]. In these systems, restoration of turbidity during reculturing requires intact cellular membranes, preserved metabolic machinery, and the ability to initiate cell division under favorable conditions. Although OD_600_ measurements do not provide absolute viable cell counts, they offer a comparative assessment of functional revival potential across treatments, particularly in carrier systems where direct enumeration may be biased by strong cell–matrix adhesion. From a practical standpoint, these results provide operational guidance for manufacturing storage-stable HAB–biochar biofertilizers: drying at 40–50 °C is preferable when immediate culturability is required, whereas drying at ≥55 °C may still be suitable for products intended for longer storage because substantial delayed recovery was observed after 30 days.

5. Conclusions

The current study shows that the post-inoculation stabilization characteristics of HAB-enriched biochar are heavily dependent upon the stabilization conditions used, especially drying temperature. The use of mild drying temperatures (40–50 °C) produced the strongest functional revival responses, preserving oxygen-containing surface functions and retaining nutrients; these results suggest that this method will produce the best conditions for creating a biologically active, slow-release nitrogen-based fertilizer. Although drying at higher temperatures (>55 °C) reduced initial culturability, some HAB populations recovered metabolic activity after 30 days of storage, suggesting stress-tolerant subpopulations and possible VBNC-like behavior or dormancy; therefore, microbial inactivation should not be equated with microbial death when developing stabilization protocols for biofertilizers. FTIR analysis showed that there were temperature-dependent losses of hydroxyl, carbonyl and C-O functional groups on the biochar surface, like the reductions observed in both ammonium binding capability and microbial attachment potential. Therefore, it can be concluded that preservation of functional group chemistry is as important as preserving microbial viability. HAB-enriched biochar has the potential to serve as a dual-function amendment that combines biological ammonification with improved nutrient retention. Future research will require evaluation of multi-strain microbial consortia, investigation of larger-scale stabilization and storage conditions, and agronomic field trial evaluations to confirm feasibility and performance under field conditions.