Next Article in Journal
Enhancing Photon Transfer Efficiency in Photocatalysis Using Suspended LED Lights for Water Treatment
Previous Article in Journal
Visible Light Induced C-H/N-H and C-X Bonds Reactions
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Reactions
Volume 4
Issue 2
10.3390/reactions4020013
reactions-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Biocatalytic Profiling of Free and Immobilized Partially Purified Alkaline Protease from an Autochthonous Bacillus aryabhattai Ab15-ES

by
Adegoke Isiaka Adetunji
1,2,3,* and
Ademola Olufolahan Olaniran
1
1
Discipline of Microbiology, School of Life Sciences, College of Agriculture, Engineering and Science, University of KwaZulu-Natal, Westville Campus, Private Bag X54001, Durban 4000, South Africa
2
Department of Biological Sciences, Summit University, Offa 250101, Nigeria
3
Centre for Mineral Biogeochemistry, University of the Free State, Bloemfontein 9031, South Africa
*
Author to whom correspondence should be addressed.
Reactions 2023, 4(2), 231-245; https://doi.org/10.3390/reactions4020013
Submission received: 1 March 2023 / Revised: 16 March 2023 / Accepted: 29 March 2023 / Published: 3 April 2023
Browse Figures
Versions Notes

Abstract

:
Partially purified alkaline protease produced by an indigenous bacterial strain, Bacillus aryabhattai Ab15-ES, was insolubilized in alginate beads using an entrapment technique. Maximum entrapped enzyme activities of 68.76% and 71.06% were recorded at optimum conditions of 2% (w/v) sodium alginate and 0.3 M calcium chloride. Biochemical profiling of free and immobilized proteases was investigated by determining their activity and stability as well as kinetic properties. Both enzyme preparations exhibited maximum activity at the optimum pH and temperature of 8.0 and 50 °C, respectively. However, in comparison to the free enzyme, the immobilized protease showed improved pH stability at 8.0–9.0 and thermal stability at 40–50 °C. In addition, the entrapped protease exhibited a higher Vmax and increased affinity to the substrate (1.65-fold) than the soluble enzyme. The immobilized protease was found to be more stable than the free enzyme, retaining 80.88% and 38.37% of its initial activity when stored at 4 °C and 25 °C, respectively, for 30 d. After repeated use seven times, the protease entrapped in alginate beads maintained 32.93% of its original activity. These findings suggest the efficacy and sustainability of the developed immobilized catalytic system for various biotechnological applications.

1. Introduction

Proteases are enzymes that catalyze the breakdown of proteins into simpler subunits by hydrolysis of the peptide bond that exists between two amino acids in a polypeptide chain [1,2,3,4]. They occupy a pivotal position, constituting more than 65% of the total industrial enzyme market [5,6,7]; their dominance of which is expected to increase further in the coming years [8]. Proteases have versatile applications in various industries, including the detergent, pharmaceutical, leather, silk, food, silver recovery and agriculture industries, as well as in wastewater treatment and organic chemical synthesis [7,9,10,11,12,13].
Proteases are ubiquitous in nature and are produced by plants, animals and microbes [14]. The inability of plant and animal proteases to meet contemporary global demands has led to an increased interest in microbial proteases. Microbial proteases serve as an excellent source of commercial enzymes since microorganisms can be cultured in large quantities in relatively short times through an established fermentation process for the generation of abundant yields of the desired metabolite. These properties are owed to their broad biochemical diversity and susceptibility to genetic manipulation [15]. Microbial proteases account for approximately 40% of the total worldwide production of enzymes [16,17,18]. Microorganisms such as bacteria, molds and yeasts have been reported for extracellular protease production [19,20].
Free enzymes are soluble and susceptible to harsh reaction conditions, including extreme temperatures, a very high or low pH, a high ionic strength, a high concentration of reactants and the presence of inhibitors [21]. Enzymes may not function optimally under such extreme conditions, which are often encountered in bioprocesses [22]. In order to ameliorate the aforementioned challenges, the immobilization of protease in suitable support materials represents a promising alternative [8,23,24,25].
The use of immobilized enzymes in bioprocesses has many advantages, including increased stability and activity over a broad range of pH and temperatures, localization, ease of product separation and handling, recovery yield and reusability, rapid termination of reactions, controlled product formation, adaptability to various engineering designs with a consequent decrease in running cost and autolysis rate [21,26,27]. However, the selection of suitable support materials and immobilization techniques is paramount to overcoming the drawbacks of immobilization. The general methods employed for enzyme immobilization include physical adsorption, ionic binding, covalent binding, cross-linking and entrapment methods [28]. Among the different immobilization techniques, the entrapment method has been considered the most often due to its simplicity and non-toxicity [29].
The entrapment method involves the physical restriction of the enzyme within a confined space or network in a semi-permeable capsule, obtained by the gelation of polyanionic or polycationic polymers in the presence of multivalent counterions, thus retaining enzyme catalytic activity and its capacity to be used continuously [8,30,31]. It is carried out in a three-dimensional gel lattice, made of either natural (agar, cellulose, alginate, carrageenan) or synthetic (polyacrylamide, polyurethane, polyvinyl, polypropylene) polymers [32,33]. However, calcium alginate beads are mostly preferred owing to their mild gelling properties, easy formulation, biocompatibility, cost-effectiveness and potential to improve enzyme stability and functional properties [34,35,36]. Alginate is a water-soluble anionic linear polysaccharide composed of sequential arrangements of 1,4-linked β-D-mannuronic acid and α-L-glucuronic acid in different proportions and can be precipitated by the addition of Ca2+ ions, producing microspheres with good strength and flexibility [37,38]. It is employed in many biotechnological applications as a support material for releasing and encapsulating cells and enzymes [39].
Proteases are classified as acidic proteases, neutral proteases or alkaline proteases based on their optimal pH of activity [14,40]. Alkaline proteases are proteases that are active at neutral to alkaline pH conditions. They occur as serine or metalloproteases and are commonly exploited for various commercial applications due to their higher activity and stability at broad pH and temperature ranges [41,42,43]. Substrates, including casein, azocasein, hemoglobin, etc., are employed for the determination of the catalytic activity of proteases. However, azocasein is mostly preferred due to its high solubility and sensitivity [44]. Protease degrades the substrate to yield trichloroacetic acid-soluble azopeptide, which is measured spectrophotometrically. Azocasein has been used as a reliable substrate for protease assays by several co-workers [45,46,47]. Many microbial species are known to produce alkaline protease. However, among bacteria, members of the genus Bacillus, including Bacillus pumilus [48], Bacillus licheniformis [49], Bacillus caseinilyticus [50], Bacillus subtilis [51,52], Bacillus megaterium [53], Bacillus coagulans [54] and Bacillus alveayuensis [7], are notable for the commercial scale production of alkaline proteases for various biotechnological applications [9,55]. Proteases from Bacillus aryabhattai have been reported [56,57]; however, to the best of our knowledge, this is the first report on a comparative study of the biochemical properties of free and immobilized proteases from Bacillus aryabhattai.
Due to the emergent market and potential applications of proteases, there is a continuing interest in bioprospecting for indigenous hyperactive proteolytic bacteria with robust catalytic activity suitable for various industrial and environmental applications. In the present study, extracellular protease from Bacillus aryabhattai Ab15-ES was partially purified and immobilized in alginate beads through entrapment. Thereafter, a comparative study of the biochemical properties of the free and immobilized proteases was carried out to ascertain their possible biotechnological applications.

2. Materials and Methods

2.1. Materials

Azocasein (used as a substrate for protease assay) and sodium alginate (used as support material for enzyme immobilization) were purchased from Sigma-Aldrich (St. Louis, MO, USA). All other reagents were of analytical grade.

2.2. Partial Purification of Protease

Protease-producing Bacillus aryabhattai Ab15-ES, previously isolated from poultry processing wastewater, was cultivated in an optimized fermentation medium consisting of (in g/L): maltose, 12.35; beef extract, 5.30; NaCl, 0.5; CaCl2.2H2O, 0.1; K2HPO4, 0.3; KH2PO4, 0.4; and MgSO4.7H2O, 0.1, with the pH adjusted to 7.8. The inoculated culture medium was then incubated at 40 °C for 48 h at 150 rpm [4]. The cell-free culture supernatant (crude protease), obtained through centrifugation at 10,000 rpm for 10 min at 4 °C, was partially purified by ammonium sulfate precipitation and dialysis. Solid ammonium sulfate was added to the crude enzyme by continuous stirring up to 60% saturation. The saturated solution was kept undisturbed at 4 °C overnight. The precipitate was collected via centrifugation at 10,000 rpm for 10 min at 4 °C, re-dissolved in a small amount (1:50) of 0.05 M Tris-HCl buffer (pH 7.2) and then dialyzed overnight against the same buffer at 4 °C.

2.3. Immobilization of Partially Purified Protease in Calcium Alginate Gel Beads

Immobilization of protease was carried out according to the entrapment method of Guleria et al. [8] with modifications to use sodium alginate as support material. Sodium alginate solution was prepared by gradually adding varying concentrations (1–5%, w/v) of sodium alginate in 0.05 M Tris-HCl buffer (pH 7.2). A two-milliliter aliquot of sodium alginate suspension was mixed with 0.5 mL of a partially purified protease. This mixture was extruded dropwise into a 5 mL CaCl2 solution (0.1–0.5 M) using a hypodermic syringe from a 5 cm height with constant stirring. The beads obtained were kept for curing at 4 °C for 1 h. Thereafter, the cured beads of about 3 mm diameter were recovered from the solution through filtration, washed 3–4 times under mild agitation with 0.05 M Tris-HCl buffer (pH 7.2) and distilled water to remove unbound enzymes and then stored at 4 °C until use [30]. A similar method was used for the preparation of control alginate beads without the enzyme.

2.4. Determination of Immobilization Efficiency

The immobilization efficiency, Y, described as the percentage of bound enzyme activity observed in the immobilization, was calculated using the equation illustrated in Equation (1) [58]:
Y   ( % ) = Observed   activity Immobilized   activity × 100
where immobilized activity was determined by measuring the total residual enzyme activity that remained in the enzyme solution after immobilization and subtracting this from the total starting activity [58].

2.5. Protease Assay

Protease activity was determined according to the modified method of Secades and Guijarro [59], using azocasein as a substrate. Enzyme solution (120 µL) was added to 480 µL of azocasein solution (consisting of 1%, w/v of azocasein in 100 mL of 0.05 M Tris-HCl buffer, pH 7.2), and the mixture was incubated at 37 °C for 30 min. The reaction was terminated by the addition of 600 µL of 10% chilled trichloroacetic acid (TCA), followed by further incubation in ice for 30 min to allow the undigested protein to precipitate. Thereafter, coagulated protein was removed from the reaction mixture through centrifugation at 10,000 rpm for 10 min at 4 °C using high-speed refrigerated centrifuge (Avanti J-26 XP, Beckman Coulter, Brea, CA, USA). The obtained supernatant (800 µL) was neutralized by addition of 200 µL of 1.8 N NaOH. The absorbance was measured at 440 nm using a UV-Vis spectrophotometer. The blank consisted of TCA–substrate solution. The activity of immobilized protease was determined using the procedure described above, except that 0.5 g of microspheres was taken for assay. Lowry method was used for the estimation of total protein using bovine serum albumin as a standard [60]. One unit (U) of enzyme activity was defined as the amount of enzyme that yielded an increase in absorbance (440 nm) of 0.01 per minute under defined assay conditions.

2.6. Characterization of Free and Immobilized Protease

2.6.1. Effect of Temperature and pH on Free and Immobilized Protease Activity

The temperature at which optimum protease activity was achieved for both free and immobilized proteases was determined by carrying out enzyme assays at various temperatures (30–80 °C) for 30 min in 0.05 M Tris-HCl buffer (pH 7.2). Protease activity was measured, as described previously. The optimum pH for free and immobilized protease activity was determined at the optimum temperature for 30 min at different pH values ranging from 3.0–11.0. The buffers used include: 0.05 M citrate buffer (pH 3.0–4.5), 0.05 M acetate buffer (pH 5.0–5.5), 0.05 M phosphate buffer (pH 6.0–6.5), 0.05 M Tris-HCl buffer (pH 7.0–9.0), and glycine–NaOH buffer (pH 9.5–11.0). Azocasein was dissolved in the respective buffers and then used to assay for protease activity, as described above. The relative protease activities as percentage (%) of maximum enzyme activity were calculated and plotted against the respective temperature or pH [30]. All experiments were carried out in triplicate.

2.6.2. Effect of Temperature and pH on Free and Immobilized Protease Stability

The thermal stability of the free and immobilized proteases was determined by pre-incubating the enzyme in 0.05 M Tris-HCl buffer (pH 8.0) at different temperatures (40, 45, 50 °C) for up to 3 h. Aliquots were taken at desired intervals and assayed for residual activity under standard assay conditions. Unheated enzyme was considered as a control (100%). The pH stability of the enzyme was assessed by pre-incubating the enzyme sample in the respective buffers (0.05 M) for up to 3 h at optimum temperature [30]. Aliquots were taken at desired intervals and assayed for residual protease activity under standard experimental conditions. All experiments were performed in triplicate.

2.6.3. Kinetics Properties of Free and Immobilized Protease

The influence of substrate (azocasein) concentrations ranging from 1 to 10 mg/mL on free and immobilized protease activity was studied by measuring enzyme activity, as previously described under standard assay conditions. Lineweaver–Burk plot was constructed by plotting reciprocals of substrate concentrations against the reciprocals of enzyme velocity. From the plot, Michaelis–Menten constant (Km) and maximum velocity (Vmax) values were determined [61].

2.6.4. Storage Stability of Free and Immobilized Protease

The storage stability of free and immobilized enzymes was determined by preserving the enzymes at 4 and 25 °C. At intervals of 5 d for up to 30 d, samples of the stored enzyme were assayed for protease activity [30]. Residual activity was calculated against the initial enzyme activity taken as 100%.

2.6.5. Reusability of Immobilized Protease

In order to assess the reusability of the protease entrapped in the calcium alginate beads, the azocasein hydrolysis reaction was repeated several times at 37 °C for 30 min. Thereafter, the beads were separated, washed with Tris-HCl buffer (pH 7.2) and used to start a cycle using fresh substrate solution [30]. Protease activity was measured after every reaction time under standard assay conditions. The relative activity was calculated by taking the enzyme activity of the freshly prepared beads in the first cycle as 100%.

2.6.6. Scanning Electron Microscopy of Beads with Immobilized Protease

The surface morphology of alginate beads containing immobilized protease was studied using ZEISS EVO LS15 scanning electron microscope.

3. Results and Discussions

3.1. Partial Purification of Protease

Bacillus aryabhattai Ab15-ES was cultivated in a previously optimized, cheap fermentation medium for extracellular protease production [4]. After centrifugation of the medium, the obtained culture supernatant, regarded as the crude enzyme, was partially purified by ammonium sulfate precipitation at a 0–60% (w/v) saturation, followed by dialysis. These techniques are useful and reliable for enzyme purification since they enable the quick and in-bulk precipitation of cellular proteins from the crude enzyme [62]. The rationale for the partial purification of the enzyme was to reduce the downstream processing costs of the biocatalyst for exploitation in large-scale biotechnological applications. The results of the purification procedures are summarized in Table 1. The partially purified protease recorded a 2.19-fold increase in specific activity in comparison to the culture supernatant and a recovery of 60.98%. Sahin et al. [25] and Zhu et al. [63] reported a 2.1-fold purification and 64% recovery, respectively, following ammonium sulfate precipitation of proteases from Bacillus subtilis M-11 and Geobacillus sp. YMTC 1049.

3.2. Immobilization of Protease in Calcium Alginate Gel Beads

Enzyme entrapment depends on sodium alginate and calcium chloride concentrations for the formation of beads with high permeability and rigidity [64]. In the present study, the effects of sodium alginate concentrations ranging from 1–5% (w/v) and 0.2 M CaCl2 were investigated on the immobilization efficiency of protease from Bacillus aryabhattai Ab15-ES. The maximum immobilization efficiency (68.76 ± 2.42%) was recorded with beads prepared from 2% sodium alginate. After this, there was a gradual decrease in the immobilization efficiency, with the lowest immobilization (22.78 ± 2.07%) obtained with 1% sodium alginate (Table 2). This is probably due to the maximum leakage of the enzyme from the beads, resulting from the larger pore sizes of the beads and less tight cross-links with calcium chloride, leading to lower immobilization efficiency [24]. This corroborates the findings of Geethanjali and Subash [64] and Anwar et al. [23], in which the maximum immobilization efficiency was recorded at 2% sodium alginate. In addition, sodium alginate concentrations of 2% have been reported by several co-workers to be suitable for enzyme immobilization [8,24,65,66,67].
Since calcium chloride serves as a cross-linking agent, its concentration affects the activity and density of immobilized biomolecules [27,68]. This also influences the mechanical strength of alginate beads as well as the efficiency of immobilized systems [69]. Therefore, the concentration of calcium chloride is essential for the stability and pore size of the beads [70]. The influence of calcium chloride concentrations (0.1–0.5 M) on the immobilization efficiency was studied at 2% sodium alginate since 2% showed the highest immobilization efficiency [30]. The highest entrapped enzyme activity was obtained from beads prepared from 0.3 M calcium chloride with an immobilization efficiency of 71.06 ± 1.30% (Table 3). This supports the findings of Geethanjali and Subash [64], in which the maximum immobilization efficiency was recorded with beads formed from 0.3 M calcium chloride.

3.3. Scanning Electron Microscopic Analysis

Scanning electron microscopy of the morphology of alginate gel beads revealed the presence of irregular pores of various dimensions on the bead surface, coupled with a stable microstructure with evenly distributed layers in the epicenter of the beads (Figure 1). The porosity of the beads helps to enhance their specific surface area and increase substrate diffusion [71].

3.4. Effect of Temperature and pH on Free and Immobilized Protease Activity

The optimum temperature of free and immobilized protease activity was determined by carrying out enzyme assays at different temperatures ranging from 30 to 80 °C. As shown in Figure 2, there was an increase in the relative activities of both the soluble and immobilized enzymes as the temperature increased from 30 to 50 °C, with the maximum activity recorded at the optimum temperature of 50 °C. Thereafter, the relative activities of both free and immobilized proteases decreased up to 80 °C. However, the entrapped protease demonstrated higher relative activity in comparison to the free enzyme, especially at high temperatures (55–80 °C). Similar results were also obtained from protease immobilized on polysulfone membranes [25] and Fenugreek β-amylase covalently attached to functionalized graphene sheets [72]. At 80 °C, the relative activity of the immobilized protease is 1.6-fold (31.01%) higher than that of the free enzyme. This clearly indicated that the support material retained the tertiary structure of the enzyme at high temperatures [23]. In other words, multi-interactions between the matrix and enzyme play a crucial role in the temperature tolerance of the immobilized enzyme [30,73].
The effect of pH on the activity of soluble and entrapped proteases was investigated under varied pH values (3.0–11.0) in different buffers at 50 °C, and the results are presented in Figure 3. The catalytic activity of the enzyme increased as the pH increased, reaching its maximum activity at pH 8.0 for both free and immobilized proteases. This suggested that the chemical coupling of the enzyme to the polymer resulted in no change in its pH profile [30]. Anwar et al. [23] and Geethanjali and Subash [64] have reported similar results on the immobilization of proteases from newly isolated Bacillus subtilis KIBGE-HAS and Labeo rohita, respectively, where no change in the pH profile of the enzyme was observed before and after entrapment in calcium alginate. This phenomenon has also been observed by other authors [74,75,76]. In addition, there was a significant increase in the relative activity of the immobilized protease at a wide pH range (8.5–11.0) in comparison to the free enzyme, an indication of its resistance to the alkaline changes in the medium [77]. Similar results have also been reported in the literature for enzymes immobilized onto/in a solid matrix [66,76,78].

3.5. Effect of Temperature and pH on Free and Immobilized Protease Stability

The thermal stability of free and immobilized proteases was investigated in the temperature range of 40–50 °C for 0–180 min. As shown in Figure 4, the protease immobilized in alginate beads demonstrated better thermal stability in comparison to the free enzyme. After incubation at 50 °C for 3 h, the immobilized enzyme retained 60.58% of its initial activity, whereas the free soluble enzyme maintained only 28.41%. The significant improvement in the thermal stability of the entrapped protease may be due to its immobilization in the support material, which protects the enzyme’s tertiary structure and prevents the conformational transition of the enzyme upon heating [30,79,80]. The enhancement of enzyme stability after immobilization in alginate beads has been reported by several authors [66,81,82,83].
The pH stability was investigated by pre-incubating the free or immobilized proteases in different buffers (pH 8.0–9.0) for 3 h at optimum temperatures. Samples were taken at 30 min intervals for the determination of the residual enzyme activity. The results revealed that the immobilized protease in alginate beads demonstrated higher stability than the free enzyme, especially at pH 8.5 and 9.0, with retentions of 69.6% and 63.53% activity, respectively, after 3 h (Figure 5). These findings can be attributed to the enzyme protection provided by the microenvironment structure and properties of the alginate beads. Similar results have been reported during the immobilization of thermostable β-glucosidase in alginate gel beads and κ-carrageenase onto magnetic iron oxide nanoparticles [66,76].

3.6. Kinetics Properties of Free and Immobilized Protease

The kinetic properties represent a key parameter for determining the efficiency of the immobilization process. In the present study, the kinetic parameters (Km and Vmax) were estimated from a Lineweaver–Burk plot using azocasein as a substrate at pH 8.0 and 50 °C. The calculated Km of free and immobilized protease were 2.023 and 1.225 mg/mL, respectively. The lower Km value of the immobilized enzyme indicates an increased affinity (1.65-fold) of the enzyme to the substrate upon immobilization in alginate gel beads (Figure 6). Furthermore, the Vmax of the immobilized protease was higher (250 U/mL) than that of the free enzyme (232.56 U/mL), suggesting a more efficient conformation of the immobilized enzyme in the solid matrix [30,84]. In addition, the Vmax/Km value of the immobilized enzyme was found to be higher than the corresponding value of the free counterpart (204.08 U/mg vs. 114.96 U/mg), an indication of the high catalytic efficiency of the immobilized enzyme [85].

3.7. Storage Stability of Free and Immobilized Protease

Storage stability represents a prominent index for the evaluation of the properties of enzymes for possible commercialization [86]. The immobilization of enzymes in a suitable support matrix is an important tool for a more rigid molecule that can resist conformational changes [72]. In essence, soluble enzymes are not stable during storage, leading to a reduction in catalytic activity [87]. In the present study, free and immobilized proteases were stored separately at 4 and 25 °C for 30 d and residual activity was determined at 5 d intervals. At 4 °C, the immobilized enzyme was found to be more stable, retaining 80.88% of its original activity after 30 d in comparison to the free enzyme, which maintained 64.6%. However, no significant loss in activity was recorded for the free enzyme for up to 5 d (Figure 7). Remarkably, while the soluble enzyme was almost completely inactivated when stored at 25 °C for 30 d, the immobilized protease still retained 38.37% of its initial activity. This improved storage stability may be due to a reduction in the denaturation of the enzyme or better structural stabilization as a result of the immobilization of the enzyme. Similar results have been reported after the immobilization of enzymes in alginate gel beads [30,66,88,89].

3.8. Reusability of Immobilized Protease

The reusability of immobilized biocatalysts is recognized as one of the most important features that influences their cost-effective application for various bioprocesses [78]. In the present study, the reuse potential of immobilized protease was investigated in several cycles for substrate hydrolysis. The immobilized protease activity decreased gradually with increasing reuse cycles. The immobilized enzyme retained 32.93% of its original activity after continuous use for up to seven cycles, indicating good operational stability of the biocatalyst [30] (Figure 8). The loss in activity may be due to repeated use, which weakens the binding strength between the matrix and the immobilized enzyme, resulting in the physical loss of the enzyme from the support material [90]. In addition, the decrease in the enzyme’s activity can be linked to the distortion of the active site due to intermittent encounters with the substrate, causing a partial or complete loss of catalytic efficiency [72]. Furthermore, the pore size of the alginate beads becomes larger after repeated use, leading to increased leakage of the enzyme and reduced biocatalytic activity [90,91]. Gupta et al. [66] reported on the reusability of immobilized β-glucosidase in alginate beads. The immobilized enzyme was reused four times with a residual activity of 17.85%. Similarly, the continuous use of protease and urease immobilized in alginate beads for up to three and five cycles, respectively, has been reported [23,64,92].

4. Conclusions

Alkaline protease from an indigenous Bacillus aryabhattai Ab15-ES was successfully immobilized in alginate gel beads at optimum conditions of 2% sodium alginate and 0.3 M calcium chloride. Both free and immobilized proteases recorded optimum activity at a pH and temperature of 8.0 and 50 °C, respectively. Entrapment in alginate gel beads considerably improved the thermal and pH stability of the enzyme, as well as its storage stability in comparison to the soluble enzyme. Furthermore, the immobilized protease exhibited an increased affinity to the substrate and a higher catalytic efficiency, as well as good operational stability for up to seven reaction cycles, suggesting its significance for sustainable and economic biosynthetic processes.

Author Contributions

Conceptualization and writing—original draft preparation, A.I.A.; funding acquisition, review and editing, A.O.O. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the National Research Foundation (NRF) of South Africa under grant number: 92803 and the APC was funded by the MDPI. Opinions expressed and conclusions arrived at are those of the authors and are not necessarily to be attributed to the NRF.

Data Availability Statement

Not applicable.

Conflicts of Interest

The authors declare that they have no conflict of interest.

References

  1. Ellaiah, P.; Srinivasulu, B.; Adinarayana, K. A review on microbial alkaline proteases. J. Sci. Ind. Res. 2002, 61, 690–704. [Google Scholar]
  2. Gupta, A.; Khare, S.K. Enhanced production and characterization of a solvent-stable protease from solvent tolerant Pseudomonas aeruginosa PseA. Enzym. Microb. Technol. 2007, 42, 11–16. [Google Scholar] [CrossRef]
  3. Shankar, S.; Rao, M.; Laxman, R.S. Purification and characterization of an alkaline protease by a new strain of Beauveria sp. Process Biochem. 2011, 46, 579–585. [Google Scholar] [CrossRef]
  4. Adetunji, A.I.; Olaniran, A.O. Statistical modelling and optimization of protease production by an autochthonous Bacillus aryabhattai Ab15-ES: A response surface methodology approach. Biocatal. Agric. Biotechnol. 2020, 24, 101528. [Google Scholar] [CrossRef]
  5. Sundararajan, S.; Kannan, C.N.; Chittibabu, S. Alkaline protease from Bacillus cereus VITSN04: Potential application as a dehairing agent. J. Biosci. Bioeng. 2011, 111, 128–133. [Google Scholar] [CrossRef]
  6. Zambare, V.; Nilegaonkar, S.; Kanekar, P. A novel extracellular protease from Pseudomonas aeruginosa MCMB-327: Enzyme production and its partial characterization. New Biotechnol. 2011, 28, 173–181. [Google Scholar] [CrossRef]
  7. Annamalai, N.; Rajeswari, M.V.; Balasubramanian, T. Extraction, purification and application of thermostable and halostable alkaline protease from Bacillus alveayuensis CAS 5 using marine wastes. Food Bioprod. Process. 2014, 92, 335–342. [Google Scholar] [CrossRef]
  8. Guleria, S.; Walia, A.; Chauhan, A.; Shirkot, C.K. Immobilization of Bacillus amyloliquefaciens SP1 and its alkaline protease in various matrices for effective hydrolysis of casein. 3 Biotech 2016, 6, 208. [Google Scholar] [CrossRef] [Green Version]
  9. Deng, A.H.; Wu, J.; Zhang, Y.; Zhang, G.Q.; Wen, T.Y. Purification and characterization of a surfactant-stable high-alkaline protease from Bacillus sp. B001. Bioresour. Technol. 2010, 101, 7100–7106. [Google Scholar] [CrossRef]
  10. Jisha, V.N.; Smitha, R.B.; Pradeep, S.; Sreedevi, S.; Unni, K.N.; Sajith, S.; Priji, P.; Josh, M.S.; Benjamin, S. Versatility of microbial proteases. Adv. Enzym. Res. 2013, 1, 39–51. [Google Scholar] [CrossRef]
  11. Sawant, R.; Nagendran, S. Protease: An enzyme with multiple industrial applications. World J. Pharm. Pharm. Sci. 2014, 3, 568–579. [Google Scholar]
  12. Herman, R.A.; Ayepa, E.; Zhang, W.-X.; Li, Z.-N.; Zhu, X.; Ackah, M.; Yuan, S.-S.; You, S.; Wang, J. Molecular modification and biotechnological applications of microbial aspartic proteases. Crit. Rev. Biotechnol. 2023. [Google Scholar] [CrossRef] [PubMed]
  13. Adetunji, A.I.; Olaniran, A.O. Treatment of lipid-rich wastewater using a mixture of free or immobilized bioemulsifier and hydrolytic enzymes from indigenous bacterial isolates. Desalination Water Treat. 2018, 132, 274–280. [Google Scholar] [CrossRef] [Green Version]
  14. Rao, M.B.; Tanksale, A.M.; Ghatge, M.S.; Deshpande, V.V. Molecular and biotechnological aspects of microbial proteases. Microbiol. Mol. Biol. Rev. 1998, 62, 597–635. [Google Scholar] [CrossRef] [Green Version]
  15. Gupta, R.; Beg, Q.K.; Khan, S.; Chauhan, B. An overview of fermentation, downstream processing and properties of microbial alkaline proteases. Appl. Microbiol. Biotechnol. 2002, 60, 381–395. [Google Scholar] [PubMed]
  16. Kumar, C.G.; Takagi, H. Microbial alkaline proteases: From a bioindustrial viewpoint. Biotechnol. Adv. 1999, 17, 561–594. [Google Scholar] [CrossRef]
  17. Haddar, A.; Bougatef, A.; Agrebi, R.; Sellami-Kamoun, A.; Nasri, M. A novel surfactant-stable alkaline serine-protease from a newly isolated Bacillus mojavensis A21: Purification and characterization. Process Biochem. 2009, 44, 29–35. [Google Scholar] [CrossRef]
  18. Raval, V.R.; Pillai, S.; Rawal, C.M.; Singh, S.P. Biochemical and structural characterization of a detergent-stable serine alkaline protease from seawater haloalkaliphilic bacteria. Process Biochem. 2014, 49, 955–962. [Google Scholar] [CrossRef]
  19. Tang, X.Y.; Wu, B.; Ying, H.J.; He, B.F. Biochemical properties and potential applications of a solvent-stable protease from a highly-yield protease producer Pseudomonas aeruginosa PT 121. Appl. Biochem. Biotechnol. 2010, 160, 1017–1031. [Google Scholar] [CrossRef]
  20. Devi, P.R.; VijayaRaghavan, P.; Vasudheven, I.; Joshua, L.; VijaKumar, M. Purification and characterization of protease from Rhizopus oligosporus. Int. J. Biol. Technol. 2011, 2, 46–49. [Google Scholar]
  21. Mugdha, A.; Usha, M. Enzymatic treatment of wastewater containing dyestuffs using different delivery systems. Sci. Rev. Chem. Commun. 2012, 2, 31–40. [Google Scholar]
  22. Karam, J.; Nicell, J.A. Potential applications of enzymes in waste treatment. J. Chem. Technol. Biotechnol. 1997, 69, 141–153. [Google Scholar] [CrossRef]
  23. Anwar, A.; Ul Qader, S.A.; Raiz, A.; Iqbal, S.; Azhar, A. Calcium alginate: A support material for immobilization of proteases from newly isolated strain of Bacillus subtilis KIBGE-HAS. World Appl. Sci. J. 2009, 7, 1281–1286. [Google Scholar]
  24. Kumar, D.; Kumar, V.; Verma, A.K.; Singh, P.; Dubey, A. Characterization and immobilization of partially purified alkaline protease extracted from rhizospheric soil bacterium, Bacillus megaterium strain EN-2 and Bacillus subtilis strain EN-3. Afr. J. Microbiol. Res. 2014, 8, 33–39. [Google Scholar]
  25. Sahin, S.; Ozmen, I.; Kir, E. Purification, immobilization and characterization of protease from local Bacillus subtilis M-11. Asia-Pac. J. Chem. Eng. 2015, 10, 241–247. [Google Scholar] [CrossRef]
  26. Massolini, G.; Calleri, E. Immobilized trypsin systems coupled on-line to separation methods: Recent developments and analytical applications. J. Sep. Sci. 2005, 28, 7–21. [Google Scholar] [CrossRef]
  27. Adetunji, A.I.; Olaniran, A.O. Immobilization and characterization of lipase from an indigenous Bacillus aryabhattai SE3-PB isolated from lipid-rich wastewater. Prep. Biochem. Biotechnol. 2018, 48, 898–905. [Google Scholar] [CrossRef]
  28. Hasirci, N.; Aksoy, S.; Tumturk, H. Activation of poly (dimer acid-co-alkyl polyamine) particles for covelent immobilization of α-amylase. React. Funct. Polym. 2006, 66, 1546–1551. [Google Scholar] [CrossRef]
  29. Gangadharan, D.; Nampoothiri, K.M.; Sivaramakrishnan, S.; Pandey, A. Immobilized bacterial α-amylase for effective hydrolysis of raw and soluble starch. Food Res. Int. 2009, 42, 436–442. [Google Scholar] [CrossRef]
  30. Adetunji, A.I. Treatment of Lipid-Rich Wastewater Using Free and Immobilized Bioemulsifier and Hydrolytic Enzymes from Indigenous Bacterial Isolates. Ph.D. Thesis, University of KwaZulu-Natal, Durban, South Africa, 2017. [Google Scholar]
  31. Bonine, B.M.; Polizelli, P.P.; Bonilla-Rodriguez, G.O. Immobilization of a plant lipase from Pachira aquatica in alginate and alginate/PVA beads. Enzym. Res. 2014, 2014, 7. [Google Scholar] [CrossRef] [Green Version]
  32. Datta, S.; Rene, C.L.; Rajaram, Y.R.S. Enzyme immobilization: An overview on techniques and support materials. 3 Biotech 2013, 3, 1–9. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Mohamad, N.R.; Marzuki, N.H.C.; Buang, N.A.; Huyop, F.; Abdul Wahab, R. An overview of technologies for immobilization of enzymes and surface analysis techniques for immobilized enzymes. Biotechnol. Biotechnol. Equip. 2015, 29, 205–220. [Google Scholar] [CrossRef] [PubMed]
  34. Tumturk, H.; Demirel, G.; Altinok, H.; Aksoy, S.; Hasirci, N. Immobilization of glucose isomerase in surface-modified alginate gel beads. J. Food Biochem. 2008, 32, 234–246. [Google Scholar] [CrossRef]
  35. Ölcer, Z.; Tanriseven, A. Co-immobilization of dextransucrase and dextranase in alginate. Process Biochem. 2010, 45, 1645–1651. [Google Scholar] [CrossRef]
  36. Jamal, F.; Qidwai, T.; Singh, D.; Pandey, P.K. Biocatalytic activity of immobilized pointed gourd (Trichosanthes dioica) peroxidase-concanavalin A complex on calcium alginate pectin gel. J. Mol. Catal. B Enzym. 2012, 74, 125–131. [Google Scholar] [CrossRef]
  37. Mahajan, R.; Gupta, V.K.; Sharma, J. Comparison and suitability of gel matrix for entrapping higher content of enzymes for commercial applications. Indian J. Pharm. Sci. 2010, 72, 223–228. [Google Scholar]
  38. Buchholz, K.; Kasche, V.; Bornscheuer, U.T. Characterization of immobilized catalyst. In Biocatalysts and Enzyme Technology; Buchholz, K., Kasche, V., Bornscheuer, U.T., Eds.; Boschstraẞe: Weinheim, Germany, 2012; pp. 411–448. [Google Scholar]
  39. Blandino, A.; Macias, M.; Cantero, D. Glucose oxidase release from calcium alginate gel capsules. Enzym. Microb. Technol. 2000, 27, 319–324. [Google Scholar] [CrossRef]
  40. Sabotic, J.; Kos, J. Microbial and fungal protease inhibitors- current and potential applications. Appl. Microbiol. Biotechnol. 2012, 93, 1351–1375. [Google Scholar] [CrossRef]
  41. Gupta, R.; Beg, Q.K.; Lorenz, P. Bacterial alkaline proteases: Molecular approaches and industrial applications. Appl. Microbiol. Biotechnol. 2002, 59, 15–32. [Google Scholar]
  42. Devi, M.K.; Banu, A.R.; Gnanaprabhal, G.R.; Pradeep, B.V.; Palaniswamy, M. Purification, characterization of alkaline protease enzyme from native isolate Aspergillus niger and its compatibility with commercial detergents. Indian J. Sci. Technol. 2008, 1, 1–6. [Google Scholar] [CrossRef]
  43. Vijayaraghavan, P.; Vincent, S.G.P. Cow dung as a novel, inexpensive substrate for the production of a halo-tolerant alkaline protease from Halomonas sp. PVI for eco-friendly applications. Biochem. Eng. J. 2012, 69, 57–60. [Google Scholar] [CrossRef]
  44. Coêlho, D.F.; Saturnino, T.P.; Fernandes, F.F.; Mazzola, P.G.; Silveira, E.; Tambourgi, E.B. Azocasein substrate for determinaXtion of proteolytic activity: Reexamining a traditional method using bromelain samples. Biomed. Res. Int. 2016, 2016, 8409183. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  45. Aissaoui, N.; Marzouki, M.N.; Abidi, F. Purification and biochemical characterization of a novel intestinal protease from Scorpaena notata. Int. J. Food Prop. 2017, 20, S2151–S2165. [Google Scholar]
  46. Iversen, S.L.; Jǿrgensen, M.H. Azocasein assay for alkaline protease in complex fermentation broth. Biotechnol. Tech. 1995, 9, 573–576. [Google Scholar] [CrossRef]
  47. Wilson, P.; Remigio, Z. Production and characterization of protease enzyme produced by a novel moderate thermophilic bacterium (EP1001) isolated from an alkaline hot spring, Zimbabwe. Afr. J. Microbiol. Res. 2012, 6, 5542–5551. [Google Scholar]
  48. Jaouadi, B.; Ellouz-Chaabouni, S.; Rhimi, M.; Bejar, S. Biochemical and molecular characterization of a detergent-stable serine alkaline protease from Bacillus pumilus CBS with high catalytic efficiency. Biochimie 2008, 90, 1291–1305. [Google Scholar] [CrossRef]
  49. Sellami-Kamoun, A.; Haddar, A.; Ali, N.E.; Ghorbel-Frikha, B.; Kanoun, S.; Nasri, M. Stability of thermostable alkaline protease from Bacillus licheniformis RP1 in commercial solid laundry detergent formulations. Microbiol. Res. 2008, 163, 299–306. [Google Scholar] [CrossRef]
  50. Mothe, T.; Sultanpuran, V.R. Production, purification and characterization of a thermotolerant alkaline serine protease from a novel species of Bacillus caseinilyticus. 3 Biotech 2016, 6, 53. [Google Scholar] [CrossRef] [Green Version]
  51. Ghafoor, A.; Hasnain, S. Purification and characterization of an extracellular protease from Bacillus subtilis EAG-2 strain isolated from ornamental plant nursery. Pol. J. Microbiol. 2010, 59, 107–112. [Google Scholar] [CrossRef]
  52. Maruthiah, T.; Esakkiraj, P.; Prabakaran, G.; Palavesam, A.; Immanuel, G. Purification and characterization of moderately halophilic alkaline serine protease from marine Bacillus subtilis AP-MSU 6. Biocatal. Agric. Biotechnol. 2013, 2, 116–119. [Google Scholar] [CrossRef]
  53. Rajkumar, R.; Jayappriyan, K.R.; Rengasamy, R. Purification and characterization of a protease produced by Bacillus megaterium RRM2: Application in detergent and dehairing industries. J. Basic Microbiol. 2011, 51, 614–624. [Google Scholar] [CrossRef]
  54. Benkiar, A.; Nadia, Z.J.; Badis, A.; Rebzani, F.; Soraya, B.T.; Rekik, H.; Naili, B.; Ferradji, F.Z.; Bejar, S.; Jaouadi, B. Biochemical and molecular characterization of a thermos- and detergent-stable alkaline serine keratinolytic protease from Bacillus circulans strain DZ100 for detergent formulations and feather-biodegradation process. Int. Biodeterior. Biodegrad. 2013, 83, 129–138. [Google Scholar] [CrossRef]
  55. Joo, H.S.; Kumar, C.G.; Park, G.C.; Paik, S.R.; Chang, C.S. Bleach-resistant alkaline protease produced by a Bacillus sp. isolated from the Korean polychaete, Periserrula leucophryna. Process Biochem. 2004, 39, 1441–1447. [Google Scholar] [CrossRef]
  56. Sharma, K.M.; Kumar, R.; Vats, S.; Gupta, A. Production, partial purification and characterization of alkaline protease from Bacillus aryabhattai K3. Int. J. Adv. Pharm. Biol. Chem. 2014, 3, 290–298. [Google Scholar]
  57. Pathak, A.P.; Rathod, M.G.; Mahabole, M.P.; Khairnar, R.S. Enhanced catalytic activity of Bacillus aryabhattai P1 protease by modulation with nanoactivator. Heliyon 2020, 6, e04053. [Google Scholar] [CrossRef]
  58. Sheldon, R.A.; van Pelt, S. Enzyme immobilization in biocatalysis: Why, what, and how. Chem. Soc. Rev. 2013, 42, 6223–6235. [Google Scholar] [CrossRef] [Green Version]
  59. Secades, P.; Guijarro, J.A. Purification and characterization of an extracellular protease from the fish pathogen Yersinia ruckeri and effect of culture conditions on production. Appl. Environ. Microbiol. 1999, 65, 3969–3975. [Google Scholar] [CrossRef] [Green Version]
  60. Lowry, O.H.; Rosebrough, N.; Farr, A.L.; Rondall, R.L. Protein measurement with the Folin-phenol reagent. J. Biol. Chem. 1951, 193, 265–273. [Google Scholar] [CrossRef]
  61. Lineweaver, H.; Burk, D. The determination of enzyme dissociation constants. J. Am. Chem. Soc. 1934, 56, 658–666. [Google Scholar] [CrossRef]
  62. Purwanto, M.G.M. The role and efficiency of ammonium sulphate precipitation in purification process of papain crude extract. Procedia Chem. 2016, 18, 127–131. [Google Scholar] [CrossRef] [Green Version]
  63. Zhu, W.; Cha, D.; Cheng, G.; Peng, Q.; Shen, P. Purification and characterization of a thermostable protease from a newly isolated Geobacillus sp. YMTC 1049. Enzym. Microb. Technol. 2007, 40, 1592–1597. [Google Scholar] [CrossRef]
  64. Geethanjali, S.; Subash, A. Optimization and immobilization of purified Labeo rohita visceral protease by entrapment method. Enzym. Res. 2013, 2013, 874050. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  65. Farag, A.M.; Hassan, M.A. Purification, characterization and immobilization of a keratinase from Aspergillus oryzae. Enzym. Microb. Technol. 2004, 34, 85–93. [Google Scholar] [CrossRef]
  66. Gupta, K.A.; Kumar, V.; Dubey, A.; Verma, A.K. Kinetic characterization and effect of immobilized thermostable β-glucosidase in alginate gel beads on sugarcane juice. ISRN Biochem. 2014, 2014, 178498. [Google Scholar]
  67. Thu, T.T.M.; Krasaekoopt, W. Encapsulation of protease from Aspergillus oryzae and lipase from Thermomyces lanuginoseus using alginate and different copolymer types. Agric. Nat. Res. 2016, 50, 155–161. [Google Scholar]
  68. Sun, J.; Liu, J.; Liu, Y.; Li, Z. Optimization of entrapping conditions of nitrifying bacteria and selection of entrapping agent. In Proceedings of the 2nd International Conference on Environmental Science and Technology (IPCBEE), Singapore, 26–28 February 2011; IACSIT Press: Chengdu, China, 2011. [Google Scholar]
  69. Konsoula, Z.; Kyriakides, M.L. Thermostable α-amylase production by Bacillus subtilis entrapped in calcium alginate gel capsules. Enzym. Microb. Technol. 2006, 39, 690–696. [Google Scholar] [CrossRef]
  70. Elibol, M.; Moreira, A.R. Production of extracellular alkaline protease by immobilization of the marine bacterium Teredinobacter turnirae. Process Biochem. 2003, 38, 1445–1450. [Google Scholar] [CrossRef]
  71. Zhan, J.F.; Jiang, S.T.; Pan, L.J. Immobilization of phospholipase A1 using polyvinyl alcohol-alginate matrix and evaluation of the effects of immobilization. Braz. J. Chem. Eng. 2013, 30, 721–728. [Google Scholar] [CrossRef] [Green Version]
  72. Srivastava, G.; Singh, K.; Talat, M.; Srivastava, O.N.; Kayastha, A.M. Functionalized graphene sheets as immobilization matrix for Fenugreek β-amylase: Enzyme kinetics and stability studies. PLoS ONE 2014, 9, e113408. [Google Scholar] [CrossRef] [Green Version]
  73. He, S.; Song, D.; Chen, M.; Cheng, H. Immobilization of lipases on magnetic collagen fibers and its applications for short-chain ester synthesis. Catalysts 2017, 7, 178. [Google Scholar] [CrossRef] [Green Version]
  74. Tükel, S.S.; Hürrem, F.; Yildirim, D.; Alptekin, O. Preparation of crosslinked enzyme aggregates (CLEA) of catalase and its characterization. J. Mol. Catal. B Enzym. 2013, 97, 252–257. [Google Scholar] [CrossRef]
  75. Saravanakumar, T.; Palvannan, T.; Kim, D.H.; Park, S.M. Optimized immobilization of peracetic acid producing recombinant acetyl xylan esterase on chitosan coated-Fe3O4 magnetic nanoparticles. Proc. Biochem. 2014, 49, 1920–1928. [Google Scholar] [CrossRef]
  76. Xiao, A.; Xu, C.; Lin, Y.; Ni, H.; Zhu, Y.; Cai, H. Preparation and characterization of κ-carrageenase immobilized onto magnetic iron oxide nanoparticles. Electr. J. Biotechnol. 2016, 19, 1–7. [Google Scholar] [CrossRef] [Green Version]
  77. Quioroga, E.; Illanes, C.O.; Ochoa, N.A.; Barberis, S. Performance improvement of araujiain, a cysteine phytoprotease by immobilization within calcium alginate beads. Process Biochem. 2011, 46, 1029–1034. [Google Scholar] [CrossRef]
  78. Ibrahim, A.S.S.; Al-Salamah, A.A.; El-Toni, A.M.; Almaary, K.S.; El-Tayeb, M.A.; Elbadawi, Y.B.; Antranikian, G. Enhancement of alkaline protease activity and stability via covalent immobilization onto hollow core-mesoporous shell silica nanospheres. Int. J. Mol. Sci. 2016, 17, 184. [Google Scholar] [CrossRef] [Green Version]
  79. Mateo, C.; Palomo, J.M.; Fernandez-Lorente, G.; Guisan, J.M.; Fernandez-Lafuente, R. Improvement of enzyme activity, stability and selectivity via immobilization techniques. Enzym. Microb. Technol. 2007, 40, 1451–1463. [Google Scholar] [CrossRef]
  80. Ranjbakhsh, E.; Bordbar, A.K.; Abbasi, M.; Khosropour, A.R.; Shams, E. Enhancement of stability and catalytic activity of immobilized lipase on silica-coated modified magnetite nanoparticles. Chem. Eng. J. 2012, 179, 272–276. [Google Scholar] [CrossRef]
  81. Ahmed, S.A.; Al-domany, R.A.; El-Shayeb, N.M.A.; Radwan, H.H.; Saleh, A. Optimization, immobilization of extracellular alkaline protease and characterization of its enzymatic properties. Res. J. Agric. Biol. Sci. 2008, 4, 434–446. [Google Scholar]
  82. Zanphorlin, L.M.; Facchini, F.D.A.; Vasconcelos, F.; Bonugli-Santos, R.C.; Rodrigues, A.; Sette, L.D.; Gomes, E.; Bonilla-Rodrigues, G.O. Production, partial characterization and immobilization in alginate beads of an alkaline protease from a new thermophilic fungus Myceliophthora sp. J. Microbiol. 2010, 48, 331–336. [Google Scholar] [CrossRef]
  83. Danial, E.N.; Hamza, A.H.; Mahmoud, R.H. Characteristics of immobilized urease on grafted alginate bead systems. Braz. Arch. Biol. Technol. 2015, 58, 147–153. [Google Scholar] [CrossRef] [Green Version]
  84. Su, R.; Shi, P.; Zhu, M.; Hong, F.; Li, D. Studies on the properties of graphene oxide-alkaline protease bio-composites. Bioresour. Technol. 2012, 115, 136–140. [Google Scholar] [CrossRef] [PubMed]
  85. Cao, S.-L.; Huang, Y.-M.; Li, X.-H.; Xu, P.; Wu, H.; Li, N.; Lou, W.-Y.; Zong, M.-H. Preparation and characterization of immobilized lipase from Pseudomonas cepacia onto magnetic cellulose nanocrystals. Sci. Rep. 2016, 6, 20420. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  86. De Queiroz, A.; Passes, E.; Alves, S.; Silva, G.; Higa, O.; Vítolo, M. Alginate-poly (vinyl alcohol) core-shell microspheres for lipase immobilization. J. Appl. Polym. Sci. 2006, 102, 1553–1560. [Google Scholar] [CrossRef]
  87. Çevik, E.; Şenel, M.; Abasiyanik, M.F. Immobilization of urease on copper chelated EC-Tribeads and reversible adsorption. Afr. J. Biotechnol. 2011, 10, 6590–6597. [Google Scholar]
  88. Yahşi, A.; Şahin, F.; Demirel, G.; Tümtürk, H. Binary immobilization of tyrosinase by using alginate gel beads and poly (acrylamide-co-acrylic acid) hydrogels. Int. J. Biol. Macromol. 2005, 36, 253–258. [Google Scholar] [CrossRef] [PubMed]
  89. Bhushan, B.; Pal, A.; Jain, V. Improved enzyme catalytic characteristics upon glutaraldehyde cross-linking of alginate entrapped xylanase isolated from Aspergillus flavus MTCC 9390. Enzym. Res. 2015, 2015, 210784. [Google Scholar] [CrossRef] [Green Version]
  90. Li, L.-J.; Xia, W.-J.; Ma, G.-P.; Chen, Y.-L.; Ma, Y.-Y. A study on the enzymatic properties and reuse of cellulase immobilized with carbon nanotubes and sodium alginate. AMB Express 2019, 9, 112. [Google Scholar] [CrossRef]
  91. Verma, M.L.; Naebe, M.; Barrow, C.J.; Puri, M. Enzyme immobilization on amino-functionalized multi-walled carbon nanaotubes: Structural and biocatalytic characterization. PLoS ONE 2013, 10, e73642. [Google Scholar]
  92. Pithawala, K.; Mishra, N.; Bahadur, A. Immobilization of urease in alginate, paraffin and lac. J. Serb. Chem. Soc. 2010, 75, 175–183. [Google Scholar] [CrossRef]
Reactions 04 00013 g001 550
Figure 1. Scanning electron microscopy of surface morphology of alginate beads.
Figure 1. Scanning electron microscopy of surface morphology of alginate beads.
Reactions 04 00013 g001
Reactions 04 00013 g002 550
Figure 2. Effect of temperature on activity of free and immobilized proteases from Bacillus aryabhattai Ab15-ES. Protease activity was measured under standard assay conditions at various temperatures (30–80 °C) at pH 7.2. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Figure 2. Effect of temperature on activity of free and immobilized proteases from Bacillus aryabhattai Ab15-ES. Protease activity was measured under standard assay conditions at various temperatures (30–80 °C) at pH 7.2. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Reactions 04 00013 g002
Reactions 04 00013 g003 550
Figure 3. Effect of pH on activity of free and immobilized proteases from Bacillus aryabhattai Ab15-ES. Enzyme activity was determined at 50 °C. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Figure 3. Effect of pH on activity of free and immobilized proteases from Bacillus aryabhattai Ab15-ES. Enzyme activity was determined at 50 °C. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Reactions 04 00013 g003
Reactions 04 00013 g004 550
Figure 4. Thermal stability of free and immobilized proteases from Bacillus aryabhattai Ab15-ES. The enzyme was pre-incubated at different temperatures (40–50 °C) for 3 h at pH 8.0, and the residual enzyme activities were estimated at regular intervals under standard assay conditions. The non-heated enzymes were taken as 100%. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Figure 4. Thermal stability of free and immobilized proteases from Bacillus aryabhattai Ab15-ES. The enzyme was pre-incubated at different temperatures (40–50 °C) for 3 h at pH 8.0, and the residual enzyme activities were estimated at regular intervals under standard assay conditions. The non-heated enzymes were taken as 100%. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Reactions 04 00013 g004
Reactions 04 00013 g005 550
Figure 5. pH stability of free and immobilized proteases from Bacillus aryabhattai Ab15-ES. The enzyme was pre-incubated at pH values 8.0–9.0 for 3 h at 50 °C, and the residual enzyme activities were estimated at regular intervals under standard assay conditions. The non-heated enzymes were taken as 100%. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Figure 5. pH stability of free and immobilized proteases from Bacillus aryabhattai Ab15-ES. The enzyme was pre-incubated at pH values 8.0–9.0 for 3 h at 50 °C, and the residual enzyme activities were estimated at regular intervals under standard assay conditions. The non-heated enzymes were taken as 100%. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Reactions 04 00013 g005
Reactions 04 00013 g006 550
Figure 6. Lineweaver–Burk plot of free (◆) and immobilized (■) proteases from Bacillus aryabhattai Ab15-ES. The enzyme activity was measured at various azocasein concentrations (1.0–10.0 mg/mL) at pH 8.0 and 50 °C. The Km and Vmax values were determined using Lineweaver–Burk plot. S: substrate concentration; V: protease-specific activity.
Figure 6. Lineweaver–Burk plot of free (◆) and immobilized (■) proteases from Bacillus aryabhattai Ab15-ES. The enzyme activity was measured at various azocasein concentrations (1.0–10.0 mg/mL) at pH 8.0 and 50 °C. The Km and Vmax values were determined using Lineweaver–Burk plot. S: substrate concentration; V: protease-specific activity.
Reactions 04 00013 g006
Reactions 04 00013 g007 550
Figure 7. Storage stability of free and immobilized proteases at 4 and 25 °C. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Figure 7. Storage stability of free and immobilized proteases at 4 and 25 °C. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Reactions 04 00013 g007
Reactions 04 00013 g008 550
Figure 8. Reusability of protease immobilized in alginate gel beads. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Figure 8. Reusability of protease immobilized in alginate gel beads. Values indicate the average of triplicate values, while the error bars represent the standard deviation.
Reactions 04 00013 g008
Table
Table 1. Summary of purification steps of protease from Bacillus aryabhattai Ab15-ES.
Table 1. Summary of purification steps of protease from Bacillus aryabhattai Ab15-ES.
Purification StepTotal Activity (U)Total Protein (mg)Specific Activity (U/mg)Recovery (%)Purification Fold
Crude enzyme24,784190.31130.231001.00
(NH4)2SO4 precipitation (60%) 15,11452.90285.7160.982.19
Table
Table 2. Immobilization efficiency of alkaline protease from Bacillus aryabhattai Ab15-ES at varying sodium alginate concentrations.
Table 2. Immobilization efficiency of alkaline protease from Bacillus aryabhattai Ab15-ES at varying sodium alginate concentrations.
Sodium Alginate Concentration (%)Immobilization Efficiency (%)
122.78 ± 2.07
268.76 ± 2.42
353.90 ± 3.86
445.35 ± 1.62
539.74 ± 0.88
All values are expressed as mean from triplicate values; ± indicates SD.
Table
Table 3. Immobilization efficiency of alkaline protease from Bacillus aryabhattai Ab15-ES at varying calcium chloride concentrations.
Table 3. Immobilization efficiency of alkaline protease from Bacillus aryabhattai Ab15-ES at varying calcium chloride concentrations.
CaCl2 Concentration (M)Immobilization Efficiency (%)
0.146.81 ± 2.41
0.262.96 ± 0.58
0.371.06 ± 1.30
0.455.92 ± 2.40
0.542.87 ± 1.78
All values are expressed in mean; ± indicates SD.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Adetunji, A.I.; Olaniran, A.O. Biocatalytic Profiling of Free and Immobilized Partially Purified Alkaline Protease from an Autochthonous Bacillus aryabhattai Ab15-ES. Reactions 2023, 4, 231-245. https://doi.org/10.3390/reactions4020013

AMA Style

Adetunji AI, Olaniran AO. Biocatalytic Profiling of Free and Immobilized Partially Purified Alkaline Protease from an Autochthonous Bacillus aryabhattai Ab15-ES. Reactions. 2023; 4(2):231-245. https://doi.org/10.3390/reactions4020013

Chicago/Turabian Style

Adetunji, Adegoke Isiaka, and Ademola Olufolahan Olaniran. 2023. "Biocatalytic Profiling of Free and Immobilized Partially Purified Alkaline Protease from an Autochthonous Bacillus aryabhattai Ab15-ES" Reactions 4, no. 2: 231-245. https://doi.org/10.3390/reactions4020013

Article Metrics

Back to TopTop