Enhanced Production of Acid Phosphatase in Bacillus subtilis: From Heterologous Expression to Optimized Fermentation Strategy

Abstract

Acid phosphatases (ACPase, EC 3.1.3.2) are hydrolytic enzymes widely distributed in both plant and animal tissues. Despite their ubiquitous presence, the production and specific activity of ACPase in these sources remain suboptimal. Consequently, the development of microbial cell factories for large-scale ACPase production has emerged as a significant research focus. In this study, we successfully expressed the phosphatase PAP2 family protein (acid phosphatase) from Acinetobacter nosocomialis 1905 in Bacillus subtilis 168. The specific activity of the crude enzyme solution was 59.60 U/mg. After purification, the enzyme activity increased to 86.62 U/mL, with a specific activity of 129.60 U/mg. Characterization of the enzyme revealed optimal activity at 45 °C and a pH of 6.0. The Km value was determined to be 0.25 mmol/L using p-nitrophenylphosphoric acid disodium salt as the substrate. Additionally, the enzyme activity was found to be enhanced by the presence of Ni²⁺. Dissolved oxygen and medium were subsequently optimized during fermentation on the basis of a commercially available 5 L bioreactor. The recombinant strain B. subtilis 168/pMA5-Acp achieved maximal volumetric enzyme activity of 136.9 U/mL after 12 h of fermentation under optimized conditions: an aeration rate of 1.142 VVM (4 lpm), an agitation speed of 350 rpm, and an optimal ratio of lactose to fish powder (7.5 g/L:12.5 g/L). These optimizations resulted in a 5.9-fold increase in volumetric enzyme activity, a 4.9-fold increase in enzyme synthesis per unit cell volume, and a 48.6% increase in biomass concentration. This study establishes a comprehensive technological framework for prokaryotic fermentation-based ACPase production, potentially addressing the bottleneck in industrial-scale applications.

1. Introduction

Elemental phosphorus is an essential macronutrient for all organisms, playing a crucial role in nucleic acid structure, membrane integrity, energy metabolism, and numerous regulatory processes [1]. The availability of phosphate in soils is often limited due to its binding to organic compounds and minerals, which restricts plant growth and yield [2,3]. Acid phosphatase (ACPase, EC 3.1.3.2) hydrolyzes phosphate in soil, enhancing its availability to plants. Moreover, ACPase regulates certain metabolic proteins, thereby promoting plant growth and development [4,5]. Consequently, the study of ACPase holds significant application potential for enhancing soil phosphorus availability and supporting plant phosphorus source regeneration.

The primary method for obtaining ACPase currently involves extraction from plant or animal cells. Recent studies have exemplified this approach, including molecular characterization and functional analysis of the tartrate-resistant acid phosphatase gene in American redfish [6]; biomimicry of purple acid phosphatase [7]; the isolation of a novel ACPase from cactus leaves [8]; the isolation, purification, and characterization of a novel ACPase from Echinacea purpurea seedlings [9]; sequence-based identification of an amyloid β-hairpin revealing a prostate ACPase fragment that promotes semen amyloid formation [10]; and the identification of acid phosphatase (ShACP) in freshwater river crabs [11].

Although microbial fermentation for phosphatase production has been relatively underexplored, research in this field has emerged over the past two decades. These studies primarily focus on microbial phosphatase production and related genetic engineering techniques, such as the process study of ACPase production via solid-state fermentation of Aspergillus niger [12] and the expression and optimization of the ACPase gene in yeast [13]. However, relatively few studies have reported on ACPase production by prokaryotic microorganisms, with most remaining at the stage of gene modification at the laboratory level. Examples include the solubilization and characterization of recombinant class C acid phosphatase from the Sphingomonas sphaericus RSMS strain [14] and the expression of a constructed acid phosphatase clone gene using Escherichia coli as the host [15].

Despite some reports on acid phosphatase production through microbial fermentation, studies on high-density fermentation in prokaryotic microorganisms for ACPase production remain scarce [16,17]. This scarcity indicates a weak foundation for research on the industrialization of this enzyme, necessitating in-depth studies. The core of this research is to control the ACPase production process through the fermentation of industrial strains, focusing on parameters such as carbon source concentration, bacterial concentration, enzyme production rates, and total enzyme activity. This approach aims to optimize the process of high-density fermentation and enzyme production.

Fermentation engineering technology plays a crucial role in efficient enzyme production. By optimizing the fermentation process, it is possible to significantly increase the cell density of production strains while enhancing enzyme expression [18,19]. This technology, situated in the midstream of the fermentation process, influences both the production capacity maximization in selected or genetically engineered strains and the efficiency of downstream product isolation [20,21]. The fermentation process is characterized by complexity, high temporal variability, and batch-to-batch variation. With the rapid advancement of industrial biotechnology, there is an increasing demand for precise control and dynamic optimization of fermentation processes. In cases where the intrinsic mechanisms of microbial metabolism are not fully elucidated, fermentation kinetics principles are employed to analyze the process through mathematical modeling [22,23]. This approach enables the accurate prediction of experimental outcomes and the optimization of the fermentation process. Kinetic modeling for fermentation control has been widely adopted by researchers and implemented in industrial applications [24,25]. In this study, we developed a kinetic model to accurately control the substrate consumption rate, growth rate, and enzyme production rate of recombinant Bacillus subtilis, aiming to maximize enzyme production efficiency during ACPase fermentation.

The current industrial production of ACPase faces significant challenges, including low yield, high costs, and complex post-processing procedures. This study aimed to develop an efficient prokaryotic expression system for ACPase production using B. subtilis 168 as the host strain. Our research focused on three main objectives: (1) establishing an efficient expression system for ACPase production; (2) optimizing fermentation conditions [26]; and (3) developing a scalable process for industrial applications. To achieve these goals, we first identified and characterized a novel ACPase gene from A. nosocomialis 1905. The recombinant strain B. subtilis 168/pMA5-Acp was then constructed for heterologous expression. Through systematic optimization of fermentation conditions in a 5 L bioreactor, including a carbon source (lactose, 7.5 g/L), a nitrogen source (fish powder, 12.5 g/L), the aeration rate (1.142 VVM), and the agitation speed (350 rpm), we significantly enhanced ACPase production. This optimized process resulted in a 5.9-fold increase in volumetric enzyme activity (reaching 136.9 U/mL) and a 48.6% increase in biomass production compared to initial conditions. Our findings demonstrate that the B. subtilis-based expression system, combined with optimized fermentation parameters, offers a promising platform for efficient and cost-effective ACPase production at an industrial scale.

2. Materials and Methods

2.1. Strains and Culture Media

A. Nosocomialis 1905, isolated from soil and capable of phosphorus degradation, was deposited in the China Center for Type Culture Collection (CCTCC) under accession number CCTCC M 20242039. Escherichia coli (E. coli) DH5α was purchased from TransGen Biotech (Beijing, China). The B. subtilis 168 and shuttle plasmid pMA5 were deposited by our laboratory.

All media were sterilized through autoclaving at 121 °C for 20 min, unless otherwise specified. The following media and solutions were prepared:

LB medium (g/L): Tryptone, 10.0; yeast extract, 5.0; NaCl, 10.0.

100× CAYE solution: Yeast extract, 10% (w/v); casein hydrolysate, 2% (w/v); autoclaving at 115 °C for 20 min.

100× EGTA solution: EGTA, 10 mM (pH adjusted to 8.0 with NaOH); autoclaving at 115 °C for 20 min.

SP I-A solution (w/v): K₂HPO₄·3H₂O, 2.8%; KH₂PO₄, 1.2%; (NH₄)₂SO₄, 0.4%; sodium citrate dihydrate, 0.2%.

SP I-B solution:MgSO₄·7H₂O, 0.04% (w/v); autoclaving at 115 °C for 20 min.

SP I medium (per 10 mL): SP I-A solution, 4.6 mL; SP I-B solution, 4.6 mL; 100× CAYE solution, 100 μL; 50% (w/v) glucose solution, 100 μL.

SP II medium (per 10 mL): SP I medium, 9.8 mL; 50 mM CaCl₂, 100 μL; 250 mM MgCl₂, 100 μL.

Seed culture medium (g/L): peptone, 10.0; NaCl, 5.0; yeast extract, 5.0; glucose, 1.0; pH 7.0 ± 0.2.

Initial fermentation medium (g/L): glucose, 5.0; yeast extract, 6.8; peptone, 2.5; MgSO₄, 0.5; NaCl, 10.0; pH 7.6.

Optimized fermentation medium (g/L): Lactose, 7.5; yeast extract, 6.8; fish powder, 12.5; MgSO₄, 0.5; NaCl, 10.0; pH 7.6.

2.2. Construction, Preparation, and Purification of Acid Phosphatase

B. subtilis 168, stored in glycerol (to a final concentration of 10%) at −40 °C, was streaked and activated on an LB solid agar plate and incubated at 37 °C for 12 h. Single colonies were picked and inoculated in 10 mL LB liquid medium, then incubated at 37 °C and 180 rpm for 12 h. An amount of 200 µL of the bacterial solution was transferred into the SP I medium and incubated at 37 °C and 180 rpm for 5 h. Subsequently, 1 mL of the bacterial solution was transferred into the SP II medium and incubated at 37 °C and 180 rpm for 1.5 h. An amount of 100 µL of 100× EGTA was then added to the SP II medium and incubated at 37 °C and 180 rpm for 10 min to obtain the receptor cells.

Acp gene amplification primers (Acp F1: aaaaggagcgatttacatatgATGAATCAACAAAAACATTTTTTTCAG and Acp R1: gagctcgactctagaggatccTTAGTGGTGGTGGTGGTGACTAAGCTCTTGTTTTAATTGTTTTTC) were designed based on the whole genome sequence of A. nosocomialis 1905. PCR amplification was performed using Phanta Max Super-Fidelity DNA Polymerase (Vazyme, Nanjing, China). The target fragment was isolated through gel electrophoresis and purified. Linear plasmid pMA5 was prepared by digestion with BamH I and Nde I, and then recombined with the purified Acp gene fragment to construct pMA5-Acp. The recombinant plasmid was transformed into competent B. subtilis 168 cells and incubated at 37 °C, 180 rpm for 90 min. Transformants were selected on LB agar containing kanamycin (50 mg/L). PCR screening of five randomly selected colonies was conducted to verify the presence of the insert. The confirmed recombinant strain B. subtilis 168/pMA5-Acp was stored at −80 °C in glycerol.

A single colony of B. subtilis 168/pMA5-Acp was picked and inoculated into the LB medium containing kanamycin (50 μg/mL). The culture was incubated overnight at 37 °C with shaking at 180 rpm. Cells were collected by centrifugation and washed with Tris-HCl buffer (50 mM, pH 7.0), and then resuspended in 5 mL of buffer. Lysozyme (1 mg/mL) was added and the mixture was incubated overnight at 4 °C, followed by sonication. The target protein was purified from the supernatant using Ni-NTA affinity chromatography and was eluted with a 2-fold column volume. A portion was analyzed by SDS-PAGE and the remainder was stored at −80 °C.

2.3. Methods of Enzyme Activity Determination

A method described by Marzadori et al. [23], including modifications, was used to test the ACPase’s activity. Using disodium p-nitrophenylphosphate (pNPP-NA2) as the substrate, pNPP-NA2 was dissolved in a 0.2 M HOAc-NaOAc pH 5.0 buffer (containing 5 mM βME) to formulate a 5 mM substrate solution. An amount of 1 mL of 5 mM pNPP-NA2, 1 mL of 50 μM MgSO₄, 3 mL of 0.2 M HAc-NaAc was preheated at 37 °C for 10 min; 100 μL of enzyme diluent was added, and was reacted at 37 °C for 10 min; 2.5 mL of 0.2 M NaOH was added to adjust the pH of the reaction solution to alkaline, so as to make it develop color, and then it was cooled down in iced water for 5 min; then, the absorbance of the reaction solution was measured. The absorbance was measured at 405 nm. For the blank control, an equal volume of the HOAc-NaOAc buffer solution was used instead of the enzyme solution, and the reaction was carried out under the same conditions. One unit (U) of acid phosphatase activity was defined as the amount of enzyme that catalyzes the release of 1 μmol of p-nitrophenol per minute under the assay conditions described above. The specific enzyme activity was defined as the number of active units per milligram of enzyme protein (U/mg). The protein content was assayed according to the instructions provided in the BCA Protein Concentration Kit [27,28].

2.4. ACPase’s Optimal Temperature and Thermal Stability

A method described by Sun et al. [29], including modifications, was used to test the ACPase’s optimal temperature and thermal stability. The enzymatic activity was measured at 5 °C intervals from 30 to 70 °C. The activity at optimal temperature was defined as 100%, and relative activities at other temperatures were calculated. For thermal stability assessment, the enzyme was incubated at each temperature for 3 h, expressed as percentages relative to the initial activity, and relative activity was calculated to examine enzyme stability across temperatures.

2.5. Optimal Reaction pH and Acid-Base Stability of ACPase

A method described by Sun et al. [29], including modifications, was used to test the optimal pH and acid-base stability of ACPase. Reaction buffers with pH values ranging from 3.5 to 8.0 at 0.5 pH unit intervals were prepared to evaluate ACPase activity. The activity at an optimal pH was defined as 100% and relative activities at other pH values were calculated to determine the optimal reaction conditions. To assess pH stability, the enzyme solution was incubated at 4 °C for 1 h at each pH value, and residual activity was measured. Residual activities were measured and expressed as percentages relative to the initial activity.

2.6. Effect of Metal Ions on ACPase Activity

The effects of metal ions on ACPase activity were investigated. The reaction system contained various metal ions (Mg²⁺, Ca²⁺, Zn²⁺,Cu²⁺,Ni²⁺, Mn²⁺, Co²⁺, and Fe³⁺) at a final concentration of 10 mM. A reaction system without metal ions served as the control group. Enzyme activity was determined under optimal reaction conditions for both groups. The activity of the control group was defined as 100%.

2.7. Determination of ACPase Kinetic Parameters

The Michaelis–Menten constant (Km) of ACPase was determined using pNPP-NA2 as the substrate at concentrations ranging from 0.2 to 1.2 mM (0.2, 0.4, 0.6, 0.8, 1.0, and 1.2 mM). Enzyme activity measurements were conducted under the optimal reaction conditions (pH 5.0, 37 °C). The Km value was calculated using the Lineweaver–Burk double-reciprocal plot method to determine the Km value of the enzyme.

2.8. Bioreactor Design

A laboratory-scale fermentation system was developed for this study (Figure 1). The system consisted of a glass bioreactor (5 L working volume) with a height-to-diameter ratio of 2:1 (H = 300 mm, D = 150 mm). The bioreactor was integrated with the following components:

• An agitation system: A dual impeller configuration comprising a Rushton turbine for mixing and a specialized paddle for foam control.
• A process control unit: Automated management of the following:

  (1)

  Substrate feeding via a peristaltic pump,

  (2)

  pH regulation,

  (3)

  Continuous culture operation,

  (4)

  Antifoam addition,

  (5)

  Agitation speed control [23].

• Monitoring and control devices:

  (1)

  A dissolved oxygen probe for real-time oxygen measurement,

  (2)

  Temperature control through a heating jacket with auxiliary water cooling,

  (3)

  An air supply system with sterile filtration.

• A data acquisition system for continuous process monitoring.

The detailed impeller configuration is shown in Figure 1.

2.9. A Dynamic Method to Determine the Volumetric Mass Transfer Coefficient K_La for a 5L Bioreactor Under Non-Culture Conditions

An amount of 3.5 L of the basic LB medium was poured into the fermenter. The fermenter was sealed and autoclaved with a dissolved oxygen electrode and antifoam agent at 121 °C for 20 min. Following sterilization, the fermenter was placed on the heating base, and all necessary components (the temperature sensor, dissolved oxygen electrode, condensate, and air vent) were connected. The stirrer was activated, and the temperature was set to 37 °C. The pressure gauge was adjusted to maintain a constant pressure of 0.2 bar (0.02 MPa, approximately 0.2 atm) in the vessel. For dissolved oxygen electrode calibration, sterile air was initially introduced into the system. Once the dissolved oxygen reading stabilized, nitrogen gas was introduced at the same flow rate. The tank pressure was maintained at 0.2 bar throughout the calibration process. The nitrogen gas flow continued until the dissolved oxygen in the medium was completely displaced. Subsequently, the gas supply was switched back to sterile air, and the dissolved oxygen concentration profile was recorded during the transition [9].

For batch fermentation, when the reactor gas–liquid phases are well mixed, the oxygen balance is given by the following equation [30]:

$${\displaystyle \frac{d{C}_L}{dt}}=OTR-OUR$$

(1)

The OTR (oxygen transfer rate) represents the amount of oxygen transferred per unit reaction volume per unit time in mol/(m³·s) or mol/(m³·h). According to the double-membrane theory, the rate-limiting step for the transfer of oxygen between the gas–liquid phases is the rate at which oxygen passes through the liquid membrane on the outside of the bubble, and thus the OTR can be expressed in terms of the volume transfer coefficient, K_La, as follows:

$$OTR=K_{L}a\cdot \left(C^{\ast }-C_L\right)$$

(2)

where K_La is the volumetric dissolved oxygen coefficient (unit: 1/s, 1/h), and C*, C_L is the oxygen concentration (unit: mol/m³).The difference between the actual dissolved oxygen value C_L and the maximum saturated oxygen concentration C* is the driving force of the oxygen transfer rate.

The difference between the actual dissolved oxygen value C_L and the maximum saturated oxygen concentration C* is the driving force of the oxygen transfer rate.

The OUR (oxygen uptake rate) indicates the amount of oxygen consumed per unit volume of fermentation broth per unit of time and is related to the specific oxygen consumption rate Q_O2 and cell concentration C_X:

$$OUR=Q_{O_2}\cdot C_X$$

(3)

2.10. Seed Culture Preparation

The B. subtilis 168/pMA5-Acp strain, stored in a −80 °C freezer, was streak-plated onto LB agar medium supplemented with kanamycin and incubated at 37 °C for 18 h. A single colony was then selected and re-streaked for isolation, followed by another 18 h incubation to obtain a pure culture. The purified strain was subsequently inoculated into the LB medium in a 250 mL Erlenmeyer flask (20% working volume) and cultured at 37 °C with shaking and 160 rpm for 16 h. Following this initial culture, 5% of the primary culture was transferred to a 1000 mL Erlenmeyer flask containing fresh LB medium (20% working volume) and incubated under the same conditions (37 °C, 160 rpm) for an additional 16–18 h.

2.11. Effect of Carbon and Nitrogen Source Type and Its Concentration on Bacterial Volume and Enzyme Production

A method described by Zhang et al. [31], including modifications, was used to ensure the optimal carbon source. Different carbon sources such as sucrose, glucose, lactose, soluble starch, and molasses (at a concentration of 10 g/L) were added to the initial fermentation medium (after the removal of glucose) and compared with each other. Other conditions were kept constant, and the effects of each carbon source on enzyme production and bacterial concentration were observed. Based on the experimental results, the carbon source with the most significant effect on the recombinant bacteria was selected, and the effect of this carbon source on enzyme production and bacterial concentration at different concentrations (2.5–12.5 g/L) was further investigated to determine the optimal carbon source concentration.

A method described by Zhang et al. [31], including modifications, was used to ensure the optimal nitrogen source. After optimization of the carbon source, different nitrogen sources such as urea, peptone, fish powder, beef extract, and ammonium sulfate (at a concentration of 5 g/L) were added to the above fermentation medium (after the removal of peptone) and compared with each other. The effect of the different nitrogen sources on enzyme production and bacterial concentration were evaluated by comparing the medium with the medium without additional nitrogen sources, keeping all other conditions constant. Based on the experimental results, the most influential nitrogen source was selected, and its effects on enzyme production and bacterial concentration were further investigated at different concentration ranges (5–15 g/L) to determine the optimal nitrogen source concentration.

2.12. The Determination of the Basal Growth Curve of the Recombinant B. subtilis 168/pMA5-Acp in a Home-Built Bioreactor

An amount of 3.5 L of basal LB liquid medium was added to 5 L of the bioreactor (η = 0.7) for sterilization. The seed solution was prepared through shaking bed oscillation, and after 16–18 h of incubation, the inoculum was inoculated into the sterilized bioreactor at an inoculum volume of 5%. The fermentation culture was carried out under the conditions of the optimal oxygen mass transfer rate obtained from the previous experiments, and the changes in bacterial concentration were characterized by OD600. After 24 h of incubation, the growth curve of the recombinant bacterium B. subtilis 168/pMA5-Acp was plotted with the fermentation time as the horizontal coordinate (h) and the OD₆₀₀ value as the vertical coordinate.

2.13. The Determination of the Enzyme Production Curve of the Strain

Samples collected at two-hour intervals were placed in centrifuge tubes and after centrifugation, the obtained organisms were stored in a refrigerator at 4 °C. After 24 h, all samples were subjected to bacterial fragmentation and centrifuged in order to obtain the fragmented supernatant for enzyme viability determination. The enzyme production curve of recombinant B. subtilis 168/pMA5-Acp was plotted using the fermentation time as the horizontal coordinate (h) and specific enzyme activity (U/mL) as the vertical coordinate.

3. Results and Discussion

3.1. Construction of High Acid Phosphatase-Producing Recombinant B. subtilis 168/pMA5-Acp

The acid phosphatase gene (Acp) was amplified from the A. nosocomialis 1905 genome, yielding a 699 bp fragment that matched the expected size (available in the Supplementary Materials (Figure S1A)). The purified Acp fragment was cloned into the shuttle vector pMA5 using a one-step cloning method and transformed into B. subtilis 168. The resulting recombinant strain B. subtilis 168/pMA5-Acp was confirmed by PCR amplification, showing the expected 699 bp band (Figure S1B). SDS-PAGE analysis of the crude enzyme solution secreted by the recombinant strain revealed a distinct band at approximately 25 kDa, corresponding to the theoretical molecular mass of 26.6 kDa (Figure S1C). The band intensity was notably higher compared to the control group, indicating the successful expression of ACPase. Notably, our expressed proteins are smaller relative to those expressed by Lu et al. [32] and Smiley-Moreno et al. [33]; proteins typically consume less energy during synthesis, folding, and degradation, which is advantageous for cellular energy conservation, and are structurally simpler and potentially more stable, reducing the risk of misfolding.

Following purification, the enzyme exhibited an activity of 86.62 U/mL, and the specific enzyme activity was 129.6 U/mg, 2.17-fold higher compared with the enzyme activity before purification (Table S1). This specific activity demonstrates a 7.63-fold enhancement compared to previously reported ACPases isolated from plant sources [9]. Moreover, our recombinant B. subtilis 168/pMA5-Acp strain showed superior enzyme production, with activities 2.25-, 2.72-, and 2.53-fold higher than those reported for Picrorhizobium-expressed ACPase using three different vector systems [34] (

Table 1. Comparison of ACPase from different sources.

Serial Number	Enzyme Activity (U/mL)	Specific Activity (U/mg)	Factor of Difference	Ref.
1	86.62	129.6	1	This study
2	-	16.98	7.63	[9]
3	38.5	-	2.25	[34]
4	31.9	-	2.72	[34]
5	34.3	-	2.53	[34]

). The recombinant B. subtilis 168/pMA5-Acp obtained from this study showed a significant lead in terms of enzyme production performance. This indicates that the recombinant strain obtained in this study has a very high potential and advantage for application in ACPase enzyme production.

3.2. Enzymatic Property Studies

To characterize the purified ACPase, we conducted systematic studies of its enzymatic properties; the ACPase exhibited the highest enzyme catalytic efficiency at 45 °C (Figure 2A), with stability maintained between 30 and 55 °C. However, prolonged exposure (3 h) to temperatures above 55 °C resulted in significant activity loss, with only 7.1% activity retained at 70 °C. Our results are closer to those of Behera et al. who found that Serratia marcescens had the highest ACPase production at 45 °C, with a decreasing trend in ACPase production above the optimal temperature [35]. However, it was also shown that Bacillus haynesii strain ACP1 had the highest ACPase activity at 50 °C [36]. Overall, these results suggest that the enzyme has poor tolerance to high temperatures, likely due to conformational changes that lead to a marked decrease in enzyme activity.

In addition to temperature effects, we also examined the influence of pH on enzyme activity and stability. As shown in Figure 2B, the enzyme showed minimal activity at an acidic pH (3.0–4.5). Activity increased with pH levels, reaching its maximum at a pH of 6.0, followed by a gradual decline at higher pH values. The enzyme maintained over 75% of its maximum activity within the pH range of 5.0–7.0. Stability studies revealed that the enzyme retained more than 90% activity between a pH of 5.0 and 7.5, indicating broad pH tolerance. However, activity decreased markedly above a pH of 7.5. The above results are the same as those of Lu et al., who found that the optimum pH for ACPase is neutral [32]. In addition, Abdelgalil et al. showed that ACPase activity was highest at a pH of 7.0 (49 U L⁻¹); at a pH of 8.0, ACPase activity decreased to 47.7 U L⁻¹ [36].

According to the experimental results shown in Figure 3, the effects of different metal ions on the enzyme activity varied significantly. Ni²⁺ showed activation with increasing concentration, and this phenomenon is similar to the findings of Lu et al. Although activation or inhibition by Zn²⁺ was not significant overall, it slightly activated the enzyme activity at low concentrations but inhibited the enzyme activity at high concentrations (4 mmol/L), but the results of Lu et al. showed that Zn²⁺ exhibited an inhibitory effect for ACPase [32]. Among the various ions tested, Fe³⁺ exhibited the most significant inhibitory effect on enzyme activity. Enzyme activity was completely suppressed when the Fe³⁺ concentration reached 2 mmol/L. Other ions such as Mg²⁺, Ca²⁺, Cu²⁺, Mn²⁺, and Co²⁺ showed minimal or negligible effects on the enzyme activity under the tested conditions. However, Lu et al. found that the inhibitory effect of Mn²⁺ and Co²⁺ on ACPase was not as significant as ours [32]. We speculate that enzymes from different sources may differ in their amino acid sequence and three-dimensional structure. These differences may lead to different conformations of the metal ion binding sites, which may affect the effect of metal ions on enzyme activity. Second, enzymes from different sources may have different dependencies on metal ions. Finally, we also performed a kinetic analysis of ACPase. Kinetic analysis of ACPase was conducted using pNPP-NA2 as the substrate. The Lineweaver–Burk plot yielded a linear equation of y = 0.2815x + 1.1369 (R² = 0.9999), from which the Michaelis constant (Km) was determined to be 0.25 mM, with a maximum reaction velocity (Vmax) of 0.88 mmol·L⁻¹·min⁻¹ (available in the Supplementary Materials (Figure S2)). The obtained Km value is marginally higher than the previously reported value of 0.167 mM [9]; however, both values remain within the same order of magnitude (10⁻¹ mM), suggesting comparable substrate affinity. The observed differences in kinetic parameters can be attributed to variations in enzyme sources, expression systems, and microenvironmental conditions. Notably, both enzymes demonstrate sufficient catalytic efficiency for industrial applications.

3.3. Effect of Different Carbon and Nitrogen Sources on Biomass and Enzyme Production During Fermentation of Recombinant B. subtilis 168/pMA5-Acp

Carbon sources are essential for cellular growth and development, significantly influencing cell density in microbial fermentation processes and indirectly affecting cellular respiratory intensity. Consequently, they are indispensable components of cultivation media. In this study, we evaluated glucose, sucrose, lactose, molasses, and soluble starch as carbon sources at a concentration of 10 g/L. We examined the optical density (OD) values of bacterial growth and relative enzyme activities under each carbon source condition using a single-factor experiment. As shown in Figure 4A, soluble starch yielded the highest relative bacterial concentration (with OD values normalized to 100% under identical experimental conditions) but resulted in lower enzyme activity. This suggests that while starch can stimulate bacterial cell proliferation, it may not favor product accumulation. Lactose, when used as the sole carbon source, resulted in the highest relative enzyme activity of the ACPase product. Molasses, as the sole carbon source, produced intermediate results for both relative bacterial concentration and enzyme activity compared to soluble starch and lactose. Given its cost-effectiveness, molasses could be considered a viable alternative for industrial cultivation media.

In this study, lactose was utilized as the carbon source in a 5 L fermentation medium. As illustrated in Figure 4B, variations in the lactose concentration significantly influenced both the growth of the recombinant strain and its enzyme activity. Lactose concentrations ranging from 2.5 to 7.5 g/L had minimal impact on bacterial growth but led to a substantial increase in the relative enzyme activity of ACPase. When lactose concentrations exceeded 7.5 g/L, a significant increase in the relative bacterial concentration was observed, accompanied by a decline in relative enzyme activity. This pattern suggested the possibility of substrate inhibition in the strain. Our findings indicated that the increase in bacterial concentration and the accumulation of product enzymes were partially coupled. Based on a comprehensive analysis of these results, we determined that 7.5 g/L was the optimal lactose concentration for the fermentation process.

The nitrogen source is a crucial component of a culture medium, primarily serving as precursor substances for microbial synthesis of amino acids, proteins, nucleic acids, and other nitrogenous metabolic products. In this study, urea, peptone, fish powder, beef extract, and ammonium sulfate were selected as nitrogen sources at a concentration of 10 g/L. Recombinant strains were cultured in fermentation media composed of these five nitrogen sources to observe bacterial growth and enzyme production (Figure 4C).

As illustrated in Figure 4C, the effect of the nitrogen source on the relative enzyme activity of the ACPase varied significantly. Ammonium sulfate and beef extract stimulated bacterial growth but resulted in low ACPase activity, with ammonium sulfate yielding the lowest enzyme activity among the tested sources. In contrast, fish powder and urea effectively increased the relative enzyme activity, suggesting their positive impact on product accumulation. To develop an accurate single-factor model for enzyme production conditions, we selected fish powder as the optimal nitrogen source for laboratory-scale fermentation in this study.

Fish powder was utilized as the nitrogen source in the laboratory-scale fermentation medium. As shown in Figure 4D, variations in the fish powder concentration significantly influenced ACPase activity. At concentrations ranging from 5.0 to 12.5 g/L, fish powder had minimal impact on bacterial growth but led to a substantial increase in relative ACPase activity. However, at concentrations exceeding 12.5 g/L, the bacterial density increase became less pronounced, and relative enzyme activity began to decline. This pattern suggested a possible substrate inhibition effect within the bacterial cells. Results from this single-factor experiment indicated that nitrogen source concentration could be partially coupled to both bacterial density increase and product enzyme accumulation. Based on a comprehensive analysis, we determined that 12.5 g/L was the optimal fish powder concentration for the fermentation process.

Our results are different from those of Behera and Abdelgalil et al., who optimally selected glucose and (NH₄)₂SO₄ as the best carbon and nitrogen sources [35,36]. We speculate that this may be related to the fermentation temperature and rotation speed, which were 37 °C and 350 rpm in our case, compared to 45 °C and 200 rpm in Behera’s case and 50 °C and 200 rpm in Abdelgalil’s case, which may have led to the differences in the strains’ utilization of carbon and nitrogen sources. In addition, different enzyme-producing strains were a factor to be considered.

The above findings suggested that optimizing the carbon and nitrogen sources in the medium significantly influenced the cell density and enzyme production of B. subtilis 168/pMA5-Acp. The experiments identified an optimal ratio of lactose to fish powder (7.5 g/L:12.5 g/L).

3.4. Effect of Aeration Ratio (Volume) and Rotational Speed on Dissolved Oxygen and OD Values During Fermentation After Media Optimization

Oxygen, as an important energy substance, is involved in the cell growth and metabolite synthesis of aerobic microorganisms; therefore, during aerobic fermentation, dissolved oxygen and its mass transfer efficiency not only determine the fermentation dynamics of aerobic microorganisms but also have an important impact on the efficiency of product synthesis [37,38]. In order to further strengthen the enzyme production process conditions of recombinant B. subtilis 168/pMA5-Acp fermentation, the enzyme production of B. subtilis 168/pMA5-Acp was enhanced by the actual aerobic fermentation measurements of dissolved oxygen (DO) dynamics associated with the rotational speed, aeration, and other physical parameters. The resulting dissolved oxygen and bacterial growth curves are shown in Figure 5, Figure 6 and Figure 7.

Initial experiments conducted at an aeration rate of 0.571 VVM (the volume of air per volume of liquid per minute) and an agitation speed of 150 rpm revealed suboptimal oxygen transfer dynamics, despite using optimized media composition. Under these conditions, the optical density (OD) of the recombinant B. subtilis 168/pMA5-Acp culture remained below 2.0 after 12 h of cultivation (Figure 5A). While increasing the agitation speed showed some positive effect on culture density, the improvement was minimal, suggesting a low oxygen uptake rate (OUR) that potentially limited enzyme production (Figure 5B). Further increases in the agitation speed, though promoting higher cell density and improved oxygen dynamics, failed to adequately enhance the mass transfer efficiency or cellular OUR (Figure 5C). These results indicate that the tested conditions (a 0.571 VVM aeration rate and a 150 rpm agitation speed) were suboptimal for efficient fermentation and enzyme production in B. subtilis 168/pMA5-Acp.

As illustrated in Figure 6A, under the conditions of 1.142 VVm (4 lpm) and 150 rpm, the OD value of B. subtilis 168/pMA5-Acp after 12 h of cultivation is higher compared to that in Figure 5A, indicating that an increased aeration rate can enhance fermentation dynamics. Figure 6B,C demonstrated that, given a stable aeration rate (VVm), increasing the agitation speed further improved the oxygen transfer dynamics during the fermentation process. The OUR of the recombinant cells was significantly improved compared to that in Figure 6A. Combined with the medium optimization process, these results further confirmed that conditions of 1.142 VVm and 350 rpm were favorable for the enzyme-producing fermentation of strain B. subtilis 168/pMA5-Acp.

Figure 7A demonstrates that although the ventilation (ratio) reached 1.714 VVm (6 lpm), the OD value of B. subtilis 168/pMA5-Acp at 150 rpm was comparable to that at the ventilation (ratio) of 1.142 VVm (4 lpm) after 12 h of culture, suggesting that the increase in the ventilation volume did not sustainably improve the OUR. Figure 7B shows that the increase in rotational speed did not extend the stabilization of the high OD values, but rather shortened the shortening of the stabilization period, which in turn potentially compromised the efficiency of enzyme production. Figure 7C indicated that while increased rotational speeds could prolong the stabilization period of high OD values, the DO profiles showed slightly inferior fermentation kinetics compared to those in Figure 6C, potentially resulting in a reduced cellular OUR in later stages and consequently affecting enzyme production efficiency.

In summary, our findings indicate that low aeration rates and rotational speeds are generally unfavorable for increasing cell concentration and product accumulation in aerobic fermentation processes. Dynamic optimization of these parameters can effectively improve the oxygen transfer rate (OTR) and oxygen uptake rate (OUR) of the process. These results are consistent with those reported by Song et al., who reported similar results using E. coli for ACPase production under more intense conditions (1.5 VVM, 600 rpm) compared to our experiment [16]. The higher aeration and agitation requirements for E. coli can be attributed to its elevated metabolic and oxygen demands, which facilitate rapid growth and efficient product formation. In contrast, B. subtilis exhibits sufficient growth at lower aeration and agitation rates due to its comparatively lower oxygen requirements. However, excessive aeration rates, while maintaining high dissolved oxygen (DO) levels, can result in shortened stabilization periods during fermentation due to accelerated cellular metabolism, potentially compromising enzyme production. Furthermore, given that the growth and metabolism of B. subtilis benefit from low shear stress conditions, we established that fermentation parameters of 1.142 VVM (4 lpm) and 350 rpm were optimal for this study.

3.5. Comparison of Recombinant B. subtilis 168/pMA5-Acp Fermentation Enzyme Production Conditions Before and After Optimization

The optimal OTR and OUR results of aerobic fermentation of recombinant B. subtilis 168/pMA5-Acp were obtained on the basis of the previous studies on the optimization of physicochemical conditions of the fermentation system. Here, the condition of 1.142 VVm and 350 rpm corresponding to maintaining a higher OUR level was adopted as the physical parameter of the fermentation device, and the fermentation medium was prepared using the optimized ratio of C:N = 1:1.6, and the volumetric enzyme activity of the crude extract could reach up to 136.9 U/mL after about 14 h of incubation, which was nearly 5.9-fold higher relative to the pre-optimization (Figure 8).

As can be seen from Figure 8, the highest volumetric enzyme activity obtained from the fermentation broth of recombinant B. subtilis 168/pMA5-Acp was 23.2 U/mL before optimization, the highest volumetric enzyme activity after optimization could be up to 136.9 U/mL, and the volumetric enzyme activity of fermentation broth per unit volume of the fermentation broth increased by about 5.9-fold, which indicates that in the same fermentation system, the enzyme amount synthesized in the cell per unit volume had a more obvious increase of 4.9-fold after optimization. The maximum OD was 4.55 before optimization, the OD could reach 6.76 after optimization, and the biomass per unit volume of fermentation also increased by 48.6%. In the present study, we found that the activity of ACPase produced by B. subtilis 168 increased more than 4-fold during the exponential growth period, which is similar to the findings of Butler et al. (about 3-fold) [17]. The above results indicated that the cell biomass and enzyme production efficiency were significantly improved through the intensive study of the laboratory pilot process of aerobic fermentation of recombinant B. subtilis 168/pMA5-Acp in this section.

4. Conclusions

In this study, we successfully expressed the ACPase-coding gene from A. nosocomialis 1905 heterologously in B. subtilis 168, generating the recombinant strain B. subtilis 168/pMA5-Acp. SDS-PAGE analysis revealed that ACPase had a relative molecular mass of 26.6 kDa, and after purification, its activity reached 86.62 U/mL. Enzymatic characterization showed that ACPase exhibited maximal activity at 45 °C and a pH of 6.0, with nickel and zinc ions acting as activators, and calcium and magnesium ions as inhibitors. Using a 5 L mechanically stirred bioreactor, we investigated the effects of agitation speed and aeration rate on dissolved oxygen dynamics and the volumetric mass transfer coefficient (KLa). The optimal physical parameters were identified as an aeration rate of 1.142 VVM and an agitation speed of 350 rpm. When combined with medium optimization (7.5 g/L lactose, 12.5 g/L fish powder), the highest volumetric enzyme activity increased to 136.9 U/mL, while cell density reached OD₆₀₀ 6.76. These results represent approximately 5.9-fold and 1.48-fold improvements in enzyme activity and biomass, respectively. Overall, the optimized fermentation system significantly enhanced ACPase production.