Skip to main content
Download PDF

Bioremoval of tannins and heavy metals using immobilized tannase and biomass of Aspergillus glaucus

Microbial Cell Factories volume 23, Article number: 209 (2024) Cite this article

Abstract

Background

The presence of inorganic pollutants and heavy metals in industrial effluents has become a serious threat and environmental issues. Fungi have a remarkable ability to exclude heavy metals from wastewater through biosorption in eco-friendly way. Tannase plays an important role in bioconversion of tannin, a major constituent of tannery effluent, to gallic acid which has great pharmaceutical applications. Therefore, the aim of the current study was to exploit the potential of tannase from Aspergillus glaucus and fungal biomass waste for the bioremediation of heavy metals and tannin.

Results

Tannase from A. glaucus was partially purified 4.8-fold by ammonium sulfate precipitation (80%). The enzyme was optimally active at pH 5.0 and 40 °C and stable at this temperature for 1 h. Tannase showed high stability at different physiological conditions, displayed about 50% of its activity at 60 °C and pH range 5.0–6.0. Immobilization of tannase was carried out using methods such. as entrapment in Na-alginate and covalent binding to chitosan. The effects of Na-alginate concentrations on the beads formation and enzyme immobilization revealed that maximum immobilization efficiency (75%) was obtained with 3% Na-alginate. A potential reusability of the immobilized enzyme was showed through keeping 70% of its relative activity up to the fourth cycle. The best bioconversion efficiency of tannic acid to gallic acid by immobilized tannase was at 40 °C with tannic acid concentration up to 50 g/l. Moreover, bioremediation of heavy metal (Cr3+, Pb2+, Cu2+, Fe3+, and Mn2+) from aqueous solution using A. glaucus biomass waste was achieved with uptake percentage of (37.20, 60.30, 55.27, 79.03 and 21.13 respectively). The biomass was successfully used repeatedly for removing Cr3+ after using desorbing agent (0.1 N HCl) for three cycles.

Conclusion

These results shed the light on the potential use of tannase from locally isolated A. glaucus in the bioremediation of industrial tanneries contained heavy metals and tannin.

Introduction

Tannery effluents having a high content of tannins, mainly polyphenols, represent a serious problem in pollution. Such pollutants can be enzymatically degraded and removed by tannases through hydrolysis of ester bonds of these polyphenols avoiding their polymerization [1, 2]. Beside effluents, tannins are also present in a variety of sources including tea, coffee, spices, as well as in various fruits and vegetables such as apricots, pomegranates, grapes, dates, eggplants, and strawberries, and in nuts like peanuts [3].

Tannin acyl hydrolase (TAH, E.C.3.1.1.20) is an enzyme that involved degrading pathway of tannin and thus has a huge industrial importance. TAH, commonly called tannase, catalyzes the hydrolysis of the central and terminal ester bonds between two aromatic rings of digallate in hydrolyzable tannins, gallic acid-based esters and other tannins, releasing gallic acid and glucose as the end products [4,5,6,7]. Fungi are a major source of valuable hydrolytic enzymes as well as economical byproducts which are needed for different modern industrial and medical applications [6, 8]. Different fungi have been reported as tannase producers such as Aspergillus species [9, 10] and Penicillium species [11].

Application of tannases reached to the food, beverages, chemical and pharmaceutical industries [12]. For instance, tannase is used in elaboration of tea powder and synthesis of gallic acid which is used in the industry of the antibiotic trimethoprim [8, 13,14,15]. Similarly, tannases have a wide applicability in laundry, detergent and leather industries as homogenizing agent for preparation of high-grade leather tannins [16,17,18]. However, the application of enzymes in industry remains limited due to the high cost of the bioprocess as well as the enzyme reusability versus stability.

The tannery industry is classified as one of the most polluting industries, generating large amounts of environmentally harmful liquids through all stages of the industrial process [19]. The presence of inorganic pollutants and heavy metals in industrial effluents particularly from electroplating, paint and tanning industries is become major environmental issues. Tannery effluents are highly complex, dark brown in color due to the presence of tanneries and have high organic content that vary according to the chemicals used. Tanneries release a large amounts of waste water containing chromium, ammonium, sulphate, dyes, and tannins. Chromium salts used in tanning process generates trivalent chromium which is toxic and carcinogenic due to its high solubility in water, permeability through biological membrane interaction with proteins and nucleic acids [20, 21]. Tannins toxicity is typically correlated with ingestion rates and absorbed tannins have the potential to cause liver and kidney necrosis. The hydrolysis of tannins by tannase is a cheap treatment to reduce the degree of toxicity of the effluents from the tannery. Similarly, industrial effluents from agrochemical industries have high levels toxic, carcinogenic and reactive pollutants such as heavy metals [22]. The discharge of industrial wastewater into water bodies will expose the environment and living entities such as plants, animals and humans to the harmful heavy metals [23]. Therefore, study of the removal of such polluting heavy metals in an environment-friendly way is urgently needed. Bio-adsorption approaches of heavy metals to the surface of the fungal biomass as biosorbents are effective environmentally friendly technology [24]. Biosorption has many advantages, such as using cheap biomass (live and dead bacteria or fungi) for removal of heavy metals and low cost treatment of large volume of effluents [25]. These heterotrophic microorganisms have the ability to form mycelial network and exploit the pollutant and use it as nutrients, which make the potential in bioremediation. Distinction of the fungal biomass as low-cost adsorbent is due to their large surface area and effective surface interaction [26]. Particularly, fungi can eliminate heavy metals by different means such as ion exchange or metal ion adsorption by the functional groups on the cell wall of the fungus. Mechanism of heavy metal biosorption starts by passive binding of fungal biomass to the metal ions and involves single or combination of different processes such as ion exchange, coordination, chelation, adsorption, complexation and electrostatic interaction [27,28,29]. Many of the fermentation processes end up by huge mass of fungal waste which in turn can be used as bio-adsorbent for the heavy metals [30, 31]. Fungi can also be easily grown in large quantities using simple fermentation methods and low-cost growth media. Therefore, the economical removal of metal ions from industrial effluents may be achieved by fungal biomass. Aftab et al. (2014) reported by Aspergillus caespitosus immobilized calcium alginate beads effectively removed lead (93% removal efficiency) from real wastewater from the paint industry [32]. Chaudhary et al., (2019) studied the effectiveness of Aspergillus fumigates in the bioremediation of tannin and chromium in tannery wastewater. They reported that A. fumigates had a maximum Cr removal efficiency of 65.1% [20].

Immobilization of enzymes via immobilization techniques such as adsorption, entrapment and covalent binding is the main strategy to improve enzyme functional properties such as activity and enzyme shelf life [33,34,35]. Currently, enzyme immobilization is preferred than using free enzymes due to long enzyme shelf life, stability under extreme conditions and multiple reuse for long periods. There are different available organic and inorganic carriers used for enzyme immobilization to ensure the highest retention of enzyme activity and stability, such as polymers, biopolymers. Alginate is a polysaccharide consists of glucuronic acid and mannuronic acid moieties, which has been successfully used for encapsulation of many biological molecules due to its biocompatibility and processivity [36]. Chitosan is a biodegradable and non-toxic natural polysaccharide which has antimicrobial potential [37]. Moreover, chitosan is considered as a suitable support for preparation of immobilized enzyme resistant to chemical degradation or metal ions disturbance.

The aim of this study is the potential use of tannase from a locally isolated fungus A. glaucus in bioremediation of tannery effluents containing tannins and/or heavy metals. Optimal physicochemical parameters affecting the activity of the partially purified tannase were investigated. Moreover, enzyme reusability in bioconversion of tannins to gallic acid as well as usability of A. glaucus biomass waste in removal of heavy metals in tannery effluents were addressed as value-added bioprocess.

Methods

Chemicals

Gallic acid methyl ester (methyl gallate) was purchased from Tokyo Chemical Industry TCI - Co., Gallic acid was obtained from oxford laboratory, tannic acid was purchased from AVI-CHEM., India, and Rhodanine was purchased from Sigma Chemical Co., U.S.A. All other chemicals used were of the highest analytical grade.

Microorganism and culture conditions

Tannase producing fungus used in this study was previously isolated from residual tea and identified as A. glaucus (GenBank accession number: MH205732) [38]. Figure 1 represents the work flow of tannase production, biomass preparation from A. glaucus and their evaluation in tannin bioconversion and heavy metal removal. For inoculum preparation; 1 ml of spore suspension of A. glaucus was inoculated in 250 ml Erlenmeyer flasks containing 25 ml of Czapek-Dox medium and incubated at 30 °C. A. glaucus was grown in previously optimized Czapex-Dox medium containing (g/l); NaNO3, 2; KCl, 0.5; MgSO4.7H2O, 0.5; KH2PO4, 1.0; FeSO4.7H2O, 0.01 and tannic acid, 10, adjusted to pH 5.5, with the moisture content of 75%. Spore suspension (1 × 106 spores/ml) were used to inoculate the production medium and then incubated at 28 °C for five days.

Fig. 1
figure 1

Workflow of tannase enzyme production and biomass production from A. glaucus

Full size image

Enzyme extraction

Crude extract was extracted from the SSF-broth to obtain the extracellular tannase by adding 50 ml of citrate buffer (0.05 M, pH 5.0) to the SSF-broth. The flask was then maintained on rotary shaker at 180 rpm and 25 °C for 1 h. The culture mixture was then squeezed using a medical gauze followed by centrifugation at 11,200 rcf, 4 °C for 15 min to separate the crude extract. The clear part was used for tannase assay [39, 40].

Tannase assay

Briefly, the reaction mixture consisted of 50 µl of diluted enzyme in 100 mM sodium-phosphate buffer pH 5.0 and 50 µl of 12.5 mM methyl gallate. The reaction mixture was incubated at 30 °C and at 600 rpm for 10 min. To stop the enzymatic reaction, 60 µl of 0.667% (w/v) methanolic rhodanine solution was added to the reaction mixture and left at 25 °C for 5 min. Then, 40 µl of 500 mM KOH was added to the reaction mixture and left for 5 min. Finally, 800 µl of distilled water was added and the mixture was incubated at 25 °C for 10 min before absorbance measurement at 520 nm. One unit of tannase activity is defined as the amount of the enzyme releasing one µmole of gallic acid per minute under assay conditions.

Estimation of protein content

The protein content of the samples was determined according to the method of Bradford [41] using bovine serum albumin as a standard.

Partial purification of tannase from A. glaucus

Ammonium sulfate precipitation

Ammonium sulfate was added to the crude enzyme to reach a saturation of 80% with constant stirring at 4 °C. Subsequently, the precipitate was then collected by centrifugation at 11,200 rcf at 4 °C for 15 min. The protein was resuspended in minimal amount of citrate buffer (0.1 M, pH 5) and dialyzed against the same buffer for 24 h at 4 °C. The enzyme activity and protein content of the fractions were measured. This precipitation process was continued up to 100% ammonium sulfate saturation.

Biochemical characterization of the tannase from A. glaucus

Effect of pH on enzyme activity and stability

To determine the optimum pH for enzyme activity, the reaction mixture was carried out within the pH range of 3.0 to 8.0 using sodium citrate buffer (0.1 M, pH 3–5) and phosphate buffer (0.1 M, pH 6–8). For pH stability studies, enzyme preparations were incubated at various pH values [3,4,5,6,7,8] for 1 h before measuring the residual enzyme activity under standard assay conditions.

Optimum temperature and thermal stability

The optimal temperature for tannase activity was determined by carrying out the enzyme reaction at different temperatures from 30 °C to 70 °C under optimal assay conditions. The thermal stability of tannase was studied by measuring the residual enzyme activity after incubating the enzyme at the same temperatures (30–70 °C) for 1 h.

Effect of metal ions on enzyme activity

To determine the effect of metal ions on the tannase activity, the enzyme solutions were incubated in the assay buffer containing 10 mM of metal salts (CaCl2, CuSO4, FeCl2, MgCl2, ZnCl2, CoCl2 and CrCl3, EDTA and β-mercaptoethanol) for 1 h at room temperature. Then the residual enzyme activity was measured under the standard assay conditions.

Immobilization of A. glaucus tannase

Immobilization of A. glaucus tannase by the entrapment method using sodium alginate was addressed [42]. Briefly, appropriate volume of partially purified enzyme was mixed with the same volume of sodium alginate solution at different concentrations (3–6%). The obtained mixture was extruded dropwise into 0.2 M CaCl2 solution with shaking. Subsequently, the formed beads were allowed to harden for 1 h, after which they were washed with 50 mM sterile citrate buffer, pH 5.0 and preserved in CaCl2 solution at 4 °C until further use.

Additionally, the partially purified tannase was covalently immobilized on chitosan, which was activated by 2.5% (v/v) glutaraldehyde [43]. Briefly, one gram of chitosan was dissolved in 100 ml of HCl containing glutaraldehyde at 30 °C for 2 h. Precipitation of the solubilized chitosan was achieved using 1 M NaOH. The precipitate was obtained by filtration and washed twice with distilled water to remove excess glutaraldehyde. Immobilization of tannase was performed by adding one ml of an enzyme solution in citrate buffer (50 mM, pH 5.0) to 5.0 mg of chitosan. The mixture was slowly stirred for 1 h at 30 °C. After that, the solids were separated, washed with the same buffer and tested for enzymatic activity. The immobilization efficiency was expressed by the ratio of specific activity of immobilized enzyme to the free one.

$$\eqalign{& Immobilization \,efficiency \left( \% \right){\rm{ = }} \cr & \left[ {{\matrix{{\rm{Specific}}\>{\rm{activity}}\>{\rm{of}} \hfill \cr {\rm{the}}\>{\rm{immobilized}}\>{\rm{enzyme}} \hfill \cr} \over \matrix{{\rm{Specific}}\>{\rm{activity}}\>{\rm{of}} \hfill \cr {\rm{the}}\>{\rm{free}}\>{\rm{enzyme}} \hfill \cr \>({\rm{non}} - {\rm{immobilized}}\>{\rm{enzyme}}) \hfill \cr} }} \right] \times 100 \cr}$$

, where the specific activity (U/mg) of the free enzyme = 5.55.

Scanning microscopy (SEM) analysis of the immobilized beads

The surface of the sodium alginate beads modified by immobilization of A. glaucus tannase was visualized using scanning microscopy technique (SEM) at 20 kV accelerating voltage. Na alginate beads was used as control. All samples were coated with gold (Sigma-Aldrich, USA) and visualized using QUANTA FEG250 (Eindhoven, The Netherlands). The magnification X500 and X2000 were used to examine the surface of the immobilized beads compared to the control. Separate areas were investigated in order to check the possible changes in the immobilized sample versus the control.

FT-IR analysis of the immobilized beads

The FT-IR analysis was performed using Vertex 70 spectroscope (Bruker, MA, USA) equipped with ATR-diamond crystal within range of 4000 –370 cm with 1 cm resolution. For samples preparation, all samples were dried in exicator for 12 h to remove the excess water. OPUS 7.0 program (bruker, MA, USA) was used to analyze the spectra.

Bioconversion of tannins by tannase

Tannase from A. glaucus was incubated in the different tannic acid solutions with different concentrations (50 g/l, 100 g/l and 200 g/l), pH 5.0 at 28 °C under shaken conditions. Gallic acid and residual tannic acid were estimated in all samples [44]. Spectrophotometric estimation of gallic acid was carried out by the method described by Sharma et al. (2000) [45]. A 300 µl of methanolic rhodanine (0.667% in methanol) was added to the standard gallic acid or suitably diluted sample followed by addition of 200 µl of 0.5 M potassium hydroxide. After incubation at 300 °C for 5 min., 4 ml distilled water was added. The absorbance was read at 520 nm after 10 min. The total tannin content of each sample was determined according to the method described by Abd-Elmotey et al. [46]. Briefly, the assay involved mixing 0.5 ml of the sample, diluted tenfold, with 2.5 ml of Folin-Ciocalteu’s phenol reagent, also diluted tenfold. After a 5-minute incubation, 2 ml of 20% (w/v) Na2CO3 was added, and the mixture was incubated for an additional hour at room temperature. Absorbance was measured at 760 nm. Percentage of tannin degradation was calculated according to the following formula:

$$\eqalign{& Percentage \,of \,tannin \,degradation{\rm{ }}\left( \% \right){\rm{ }} = \cr & \left[ {{{{\rm{Tannin}}\>{\rm{before}}\>{\rm{degradation}} - \>{\rm{Tannin}}\>{\rm{after}}\>{\rm{degradation}}} \over {{\rm{Tannin}}\>{\rm{before}}\>{\rm{degradation}}}}} \right] \times \>100 \cr}$$

Evaluation of A. glaucus biomass waste in bioremediation of some heavy metal ions from aqueous solution

Heavy metal ions uptake by A. glaucus biomass

All bio-removal experiments were performed as batch experiments in which five grams of live fungal biomass (as a bio-removal agent) was added into 250 ml Erlenmeyer flasks individually containing 50 ml of metal ion solution (Cr3+, Pb2+, Cu2+, Fe3+ and Mn2+) and incubated in a shaking incubator at 150 rpm for 10 min. at 30 °C [47]. Similarly, 50 ml of metal(s) solution was incubated without biomass as negative control. Stock metal solutions were prepared as 1000 ppm using chloride salts; CrCl3, PbCl2, CuCl2, FeCl3 and MnCl2 (Merck (NJ, USA) or VWR chemicals BDH (PA, USA)). The biomass was harvested by filtration on filter paper (Whatman No. 1). The residual metal concentration in filtrate was analyzed for the determination of residual metal(s) by Flame Atomic Absorption Spectrophotometer (Agilent Technologies 200 series AA). For sample digestion, 10 ml of HNO3 (1:1) was mixed with the sample. The slurry was then heated at 95 °C and refluxed for 10 min. Sorption of metal ions by the biomass (metal ion uptake %) was calculated using the following expression;

$$\:Unsorbed\:metal\:\left(residual\:metal\:\%\right)=\left[\raisebox{1ex}{$\left(\text{A}\times\:100\right)$}\!\left/\:\!\raisebox{-1ex}{$\text{B}$}\right.\right]$$
$$\:Sorbed\:metal\:\%=\left[\raisebox{1ex}{$\left(100-A\right)\times\:\:100$}\!\left/\:\!\raisebox{-1ex}{$\text{B}$}\right.\right]$$

A = Metal ion concentration in supernatant.

B = Initial metal ion concentration (control).

Effect of live and dead fungal biomass on Cr3+ uptake

Fungal biomass, including both live and dead forms, was assessed for its Cr3 + uptake capacity. Accordingly, five grams of wet fungal biomass, equivalent to 0.44 g dry weight, were subjected to pre-treatment before being incubated with a Cr3 + solution (concentration of 30 ppm at pH 5) under two separate conditions. The first treatment was boiling of fungal biomass in distilled water for 10 min the second one was soaking of fungal biomass in 5% KOH solution for 10 min. The fungal biomass was then separated by filtration, washed with 1 N HCl and thoroughly washed several times with distilled water until neutralization [47]. The living fungal biomass (untreated) was served as control.

Cr3+ desorption and reloading of fungal biomass

Sorption of Cr3+ was carried out as described previously. After initial Cr3+ uptake, the fungal biomass was separated by filtration and repeatedly washed with distilled water. The metal ions content in the supernatant was determined. Sorbed metal ion by the biomass was taken as 100% for the subsequent uptake cycle. The biomass obtained from the initial biosorption (described above) was suspended in 50 ml of the desorbing agent (0.1 N HCl) for 1 h. The biomass was separated by filtration, washed and re-suspended in Cr3+ solution for 1 h. Similarly, 50 ml of the desorbing solution without the fungal biomass was considered as a negative control. By the end of incubation period, the biomass was separated and Cr3+ content in the clear filtrate was determined. The percentage of elution efficiency of the desorbing agent was calculated as follows:

$$\eqalign{& Elution \,efficiency{\rm{ }}\% {\rm{ }} = \cr & \left[ {{{{\rm{amount}}\>{\rm{of}}\>{\rm{desorbed}}\>{\rm{metal}}\>{\rm{ion}}} \over {{\rm{amount}}\>{\rm{of}}\>{\rm{biomass}}\>{\rm{used}}\>{\rm{for}}\>{\rm{metal}}\>{\rm{ion}}\>{\rm{sorption}}}}} \right] \times \>10 \cr}$$

Statistical analysis

All experiments were performed in triplicates and results were presented as mean values ± standard error (SE).

Results

Partial purification of tannase from A. glaucus

Tannase from A. glaucus was partially purified 4.83-fold with a yield of 66.67% by fractional precipitation by ammonium sulfate (80%). The specific activity of the enzyme showed substantial increase from 4.09 to 19.76 U/mg protein.

Biochemical characterization of tannase

Effect of pH on the activity and stability of tannase

The optimal pH for tannase activity was determined by measuring the enzyme activity at various pH values. As shown in Fig. (2a), the enzyme was most active at the pH range 4.0–6.0 with an optimal activity at pH 5.0. On the other hand, a sharp decrease in enzyme activity was observed above pH 6.0 indicating the acidic nature of the enzyme. Concerning pH stability of tannase, results showed that the enzyme was completely stable at pH range from 5.0 to 6.0 since no loss of enzyme activity was observed after 12 h of incubation (Fig. 2b). On the other hand, the enzyme was more stable at low pH values than at high pH values retaining about 38% of the maximum activity at pH 3.0, while the enzyme completely lost its activity at pH 8.0.

Fig. 2
figure 2

Effect of pH on the enzyme activity (a) and stability (b) of the tannase

Full size image

Effect of temperature on the activity and stability of tannase

Studies on the effect of temperature on the activity and stability of tannase revealed that the highest activity was recorded at the temperature range 30 –40 °C (Fig. 3). In addition, the thermal stability of tannase indicated that the enzyme was completely stable up to 40 °C, since no loss in catalytic activity occurred after 1 h of incubation (Fig. 4). Moreover, the enzyme still retained about 50% of its original activity after incubation at 60 °C for 1 h.

Fig. 3
figure 3

The effect of temperature on the tannase activity

Full size image
Fig. 4
figure 4

Thermal stability of tannase enzyme

Full size image

Effect of metal ions and other chemicals on tannase activity

A study on the effect of different metal ions on tannase activity indicated that the enzyme was active in the presence of all metal ions tested except Fe2+, Cu2+ and EDTA (Table 1). On the other hand, β-mercaptoethanol decreased the enzyme activity the most due to disruption of the formation of disulfide bonds.

Table 1 Effect of metal ions on tannase activity
Full size table

Immobilization of A. glaucus tannase

The partially purified tannase was immobilized by two different methods, covalent binding using chitosan and Na-alginate entrapment method. Results showed that, the enzyme immobilized on Na-alginate exhibited higher specific activity in comparison with the enzyme immobilized on chitosan. Whereas the immobilization efficiencies of A. glaucus tannase immobilized on Na-alginate and chitosan were found to be 75% and 57% respectively (Table 2). In addition, Entrapment of tannase in Na-alginate resulted in 3–4 mm compact beads. It was observed that, the concentration of Na-alginate in the initial solution affect the bead properties. The beads prepared with 3% Na-alginate were fragile and loosely formed while those prepared with 4, 5 and 6% Na-alginate were non-fragile and compact beads. In addition, the enzyme beads prepared with 3% and 4% Na-alginate concentration showed higher activity (Sp. activity 4.16 and 3.84 U/mg, respectively) than enzyme beads of 5% and 6% Na-alginate (Sp. activity 1.28 and 0.32 U/mg, respectively). While a higher polymer concentration in beads can enhance stability, it may also restrict enzyme diffusion, leading to a reduced reaction rate. The better immobilization system was chosen according to the better enzyme stability, reusability and rate of catalysis. Therefore, 3% Na-alginate was finalized as the best immobilization efficiency comparing to other matrices.

Table 2 Immobilization of A. glaucus tannase
Full size table

Examination of the surface of na alginate beads carrying tannase using the SEM Microscopy

The study of the surface of different materials used for the immobilization of enzymes requires modern tools such as scanned electron microscope (SEM).

The microscopic analysis of the surface of Na alginate beads (3%) showed morphological changes among different analyzed samples (Fig. 5). Na alginate beads was used as control (Fig. 5a) while the sample of the Na alginate carrying the A. glaucus tannase (Fig. 5b). Interestingly, the SEM micrographs revealed that the immobilization of tannase into the composition of the Na alginate beads resulted in smoothing of the biochemical matrix of the beads compared to the control. This observation suggests an enhancement of the mechanical properties of the immobilized sample.

Fig. 5
figure 5

SEM micrographs of the surface of 3% Na alginate beads (control) (Upper), SEM micrographs of the surface of 3% Na alginate beads containing tannase enzyme isolated from A. glaucus (Lower)

Full size image

Examination of the surface of na alginate beads carrying tannase using FT-IR

Infrared Spectroscopy was performed for both free Na alginate beads and beads containing tannase enzyme isolated from A. glaucus to detect the appearance of certain chemical bonds. The obtained spectra for both cases are presented in Fig. 6.

Fig. 6
figure 6

FT-IR spectra of 3% Na alginate beads (control) (Upper), FT-IR spectra of 3% Na alginate beads containing tannase enzyme isolated from A. glaucus (Lower)

Full size image

Sodium alginate beads (3%) exhibited absorption bands at 3252.06 cm− 1 which can be due to presence of hydroxyl group (-OH), 1591.25 cm− 1 and 1415.29 cm− 1 which may refer to asymmetric and symmetric stretching vibration of COO groups, respectively. Another absorption band was shown at 1080.54 cm− 1 which may represent the elongation of C-O groups (Fig. 6a). The sample of Na alginate carrying the partially purified tannase from A. glaucus showed absorption bands at 3341.08, 1120.5, 1079.75 and 605.72 cm− 1. These results indicate the existing of both O-H stretch rates in the intermolecular hydrogen bonds pattern, C-C and C-O stretches in the benzene structure bonds.

Reusability of immobilized tannase on Na-alginate

To evaluate the reusability of immobilized A. glaucus tannase, seven cycles were carried out by repeating use the Na-alginate beads after washing it well after each reaction. As shown in Fig. 7, the immobilized enzyme kept 70% of its initial activity after four reuses. The activity of immobilized enzyme was decreased by about 12% after each consecutive use till the five cycles.

Fig. 7
figure 7

Reusability test of the immobilized A. glaucus tannase at repeated enzyme reactions

Full size image

Bioconversion of tannins to gallic acid by immobilized A. glaucus tannase

The efficiency of immobilized A. glaucus tannase for conversion of tannic acid to gallic acid was studied at two different temperatures (30 °C and 40 °C). The residual tannic acid and gallic acid produced were estimated in all samples. Results showed that the efficiency of immobilized enzyme for bioconversion of tannic acid to gallic acid was higher at 40 °C than that at 30 °C (Table 3). In addition, the highest production of gallic acid (44.80 mg/ml) was obtained at 40 °C when the tannic acid concentration was 50 g/l. Moreover, the immobilized tannase can be reused for more than three cycles in the bioconversion process with the same efficiency (data not shown).

Table 3 Bioconversion of tannins by immobilized A. glaucus tannase
Full size table

Bioremediation of heavy metal ions usingA. glaucusbiomass

The utilization of A. glaucus biomass as a biosorbent for the bioremediation of heavy metal ions, Cr3+, Pb2+, Cu2+, Fe3+ and Mn2+, was investigated in order to maximize the benefit of the bioprocess. Therefore, the fungal waste biomass was used to remove some heavy metals from aqueous solution. As shown in Fig. 8, the use of fungal biomass resulted in the bioremediation of heavy metal ions from aqueous solution (30 ppm) with the remediation percentage of 37.20, 60.30, 55.27, 79.03 and 21.13 for Cr3+, Pb2+, Cu2+, Fe3+, and Mn2+ respectively.

Fig. 8
figure 8

Metal ions uptake by A. glaucus biomass

Full size image

Effect of live and dead fungal biomass on Cr3+ uptake

Since the previous biosorption experiments were based on the living fugal biomass, thus, the impacts of using dead fungal biomass (KOH-treated and boiled biomass) compared to the living biomass, as control, on the biosorption efficiency of Cr3+ was investigated. Results revealed that the biosorption process of Cr3+ was higher when the living fungal biomass was used, indicating that fungal biomass’s uptake was dependent on cellular activity. The Cr3+ uptake percentage using living biomass was 37% which exceeded that of KOH-treated and boiled biomasses (26.03% and 22.60%, respectively) (Fig. 9).

Fig. 9
figure 9

Cr3+uptake by A. glaucus biomass. Living biomass ‘control’ and dead biomass (by boiling and by KOH-treatment

Full size image

Cr3+ desorption and reloading of fungal biomass

The experiment was designed to recover the sorbed metal ions from fungal biomass by desorbing agent and reuse of the regenerated biomass in metal ions uptake. An efficient Cr3+ desorption using 0.1 N HCl as desorbing agent was noticed. Since the initial loaded Cr3+ on the fungal biomass was 11.15 ppm and the recovered Cr3+ from the fungal biomass was 5.29 ppm, thus, Cr3+ desorption using 0.1 N HCl was expressed as 47.44% Cr3+ uptake. This process was repeated for three cycles where the uptake of Cr3+ was decreased from 1st Cycle to 3rd cycle by (26, 13.60, 2.26%, respectively).

Discussion

The main objective of this study was to exploit the potential of tannase from A. glaucus in removing tannins and fungal biomass in the bio-removal of heavy metals for treating industrial effluents like those from tanneries. Partial purification of tannase from A. glaucus was achieved by ammonium sulfate precipitation at saturation of 80%. The purification resulted in approximately five folds purification relative to the crude protein and substantial increase in the specific activity (from 4.1 to 19.76 U/mg). Partially purified tannase from A. glaucus with 66.67% yield which was higher than that reported for tannases purified from Aspergillus niger with 18.40% yield [15]. In addition, fractional precipitation of the Paecilomyces variotii culture filtrate with ammonium sulfate resulted in 78.7% yield [48]. A significant reduction in the tannic acid content of beer and orange juice was achieved by tannase purified from Klebsiella pneumoniae with a 9.5% yield. Indicating that tannase may be used to enhance the quality of fruit juices and beverages [2].

The biochemical studies were investigated to reveal some of the physiochemical properties of tannase that might be significant in its industrial applications. The characterization studies indicated that optimal activity of tannase from A. glaucus was recorded by acidic pH values (4.0–6.0) with maximum activity at pH 5.0. Similarly, it was previously reported that tannase of fungal origin is mostly active at acidic conditions (pH 4.0–6.0) while tannase of bacterial origin is primarily active at wider pH range from 4.0 to 7.0 [49, 50]. The impact of the pH on the enzyme depends on the ionization state of the amino acids on the active site. It is happened through the conformational adaptation resulted from the ionization of different amino acids [51]. Moreover, the activity of tannase purified from A. glaucus was favored by acidic pH, this may be because its active sites are rich in acidic amino acids. The amino acid properties of the active site have been reported to determine pH-dependent enzyme activity [52]. Accordingly, Govindarajan, R. K., et al. reported that acidic pH (pH 6.0) has a great value for tannase application in tanneries effluent degradation and pharmaceutical industry [53]. Thus, our results are found to be similar to earlier reports on tannases from different fungi such as Aspergillus ruber and Verticillium sp P9 [54, 55].

Studies on the effect of temperature on the activity and stability of A. glaucus tannase revealed that, the enzyme was optimally active at 40 °C. A gradual decrease in enzyme activity was observed at higher temperatures. High temperature could lead to the change of secondary and tertiary structures of tannase and the enzyme stability would decrease when the conformation of the protein was destroyed. In addition, The α-helix content of tannase decreased and the ratio of randomness increased at high temperatures [56]. This findings might refer to the fact that high temperatures increases the reaction velocity and also the rate of enzyme denaturation [57]. Similarly, optimal tannases activity obtained from Penicillium notatum NCIM 923 was reported at 40 °C [58].

Concerning temperature stability, it was found that tannase from A. glaucus was stable in the temperature range 30–60 °C retaining approximately 50% of its initial activity at 60 °C. Additionally, tannase from A. glaucus exhibited a maximal ionic stability at pH range 4.0 to 6.0 as more than 90% of the maximal activity remained after 12 h of incubation at pH 5.0. These results indicate that A. glaucus tannase is a stable enzyme under different physiological conditions which is considered as an added value for tannase-assisted bioprocesses performed at high temperatures. In this context, the optimal activity and stability of tannases of different origins were reported in a range of 30–60 °C, whereas fungal tannases showed higher activity and stability under different temperatures than those from bacteria and yeast [49, 59, 60].

A study investigating the effect of different metal ions on tannase activity indicated that all the tested metal ions did not impact enzyme activity, except for Fe2+ and Cu2+, which negatively affected tannase activity. Additionally, tannase from A. glaucus was partially inhibited by EDTA which indicates that the active site of the enzyme has amino acids contains sulfur, as it was previously reported by Da costa et al. [59]. In a similar fashion, Chhokar, Vinod, et al. reported that the purified tannase from Aspergillus heteromorphus MTCC 8818 was inhibited by several metals such as Hg2+ and Ag+ including also Fe2+ and Cu2+ [61].

In this study, the immobilization technique was selected based on the immobilization efficiency and stability of the immobilized enzyme. Immobilization of tannase was achieved using different concentrations of Na alginate compared to chitosan as immobilization carriers. The results indicated that the 3% Na alginate was the best carrier for tannase immobilization among others with an immobilization efficiency of 75%. Lower immobilization efficiency of tannase was noticed by increasing the Na alginate concentrations above 3% which could be interpreted by lack of uniform pore size due to the high viscosity of the tannase-alginate mixture, as it was similarly reported by Keerti et al. [62]. Increased Na alginate concentrations in beads have the potential to improve stability, but they may also limit the diffusion of enzymes, which would lower the reaction rate. Aharwar and Parihar (2023) reported a decrease in tannase activity of Talaromyces verruculosus immobilized in alginate, may be due to the active site blockage [63]. Na alginate proved to be the best support to immobilize A. glaucus tannase, with an immobilization efficiency of 75%, which was higher than that of immobilized Aspergillus niger tannase (36.8%) [64] and Penicillium variable tannase with the immobilization yields of 69% [65]. Yao et al., (2014) found that tannase encapsulation in calcium alginate was the best immobilization method with 62%, while Amberlite IRC 50 was the least efficient support (5%, immobilization yield) [66].

Scanned electron microscopy and FT-IR spectroscopy were used as modern and precise tools to study the structure of many samples used in biological systems [67]. Other papers similarly examined the absorption spectra of the purified tannase. For instance, Govindarajan et al., 2019 have been reported similar absorption bands for their purified tannase at 3440.91, 1466.68, 1054.60 and 699.07 cm− 1 by which they indicated the ability of tannase isolated from Bacillus subtilis KMS2-2 to degrade tannic acid as substrate into simple ester molecule [53].

The enzyme reuse is the main advantage obtained from the immobilization process. The possibility of reusing the immobilized tannase-alginate preparation was assessed for seven cycles. The results revealed that the immobilized tannase retained about 70% of its initial activity after the fourth cycles, whereas the activity of immobilized enzyme was then decreased by the number of enzyme reuse. This result can be interpreted by the enzyme inactivation occurred upon the enzyme loss from the gel due to diffusion effects as well as the multiple use and of enzyme [68]. The operational stability of the immobilized tannases is one of the most important factors in industrial applications [66]. Srivastava A, Kar R. reported that Aspergillus niger tannase immobilized on Na-alginate can hydrolyze 68.8% of tannins in fresh aonla juice in the first run, but can only hydrolyze 37.7% of tannins after the second run [69]. Also, tannase immobilized on Ca alginate beads had good operational stability when it hydrolyzes 56% of tea tannins from a green tea infusion after six cycles [66].

One of the important role of the tannase enzyme is obtaining gallic acid through degradation of tannins from environmentally hazardous tanneries [1]. Gallic acid formed can be directly used as a feed additive and pharmaceutical applications [70]. The efficiency of immobilized tannase of A. glaucus for bioconversion of tannins and production of gallic acid was higher at 40 °C than 30 °C. The high enzyme activity remarked at 40 °C can be interpreted by incidence of higher substrate solubility at higher temperatures which in turn accelerate the reaction speed. On the other hand, it was found that the efficiency of bioconversion of tannic acid to gallic acid was inversely proportional to the tannic acid concentrations. This finding can be explained by lowering of the mass transfer and substrate diffusion at high tannic acid concentration [46]. Our findings demonstrated that the immobilized tannase can be used more than three cycles for bioconversion of tannic acid to gallic acid with high efficiency compared to the free tannase from A. glaucus [38].

Recently, different microorganisms showed a high capability of heavy metal biosorption from wastewater [71, 72]. Therefore, we further investigated the biosorption of heavy metals such as Cr3+ using living and dead A. glaucus biomass. To obtain dead biomass of A. glaucus, cells were boiled in first treatment and treated with KOH as alkaline sanitizer in the second treatment. The high pH of KOH (higher than 12.0) results in cell membrane lysis and final cell lysis. Results showed a higher Cr3+ uptake capacity by the living fungal biomass than that obtained using dead fungal biomas. This can be explained by the fact that heavy metal biosorption including Cr3+ uptake by fungal biomass was dependent on cellular metabolism. Passive metal uptake, at which the metal ion sorption is non-metabolic, can happen whether cells are living or dead. In contrast, active biosorption in which the heavy metals passed through the cell membrane to the living cells requires metabolic activity of the living cells [73,74,75]. Live fungal biomass also has a greater adsorption capacity of heavy metals than the equivalent dry biomass due to the abundant charges on their surfaces. The biosorption process of fungal biomass might be greater, less or equivalent than that of their living counterparts [76]. In this concern, live mass of Aspergillus sp. had a greater biosorption of Cu 2+ than dead biomass [77].

This observation was in-line with former studies reported different constituents and functional groups in the fungal cell wall which are able to attract heavy metals where the ionic state of the biomass determines the effectiveness of the biosorption [78,79,80]. The highest uptake of Cr3+ by A. glaucus biomass is pointing out the presence binding sites on the fungal cell wall which act as active biosorbent for Cr3+. The applicability of the metal biosorption process is improved by (1) potential recovery of the sorbed metal and (2) potential reuse of the biomass in multiple sorption-desorption cycles. Thus, the regenerated biomass of A. glaucus was used for three sorption-desorption cycles of Cr3+. Sukumar, C., et al. reported that reuse of adsorbed metals is highly needed to diminish the metal contamination which can be evaluated by its adsorption efficiency in multiple adsorption- desorption process [81].

These findings shed the light on the potential use of tannase from A. glaucus in biodegradation of hydrolysable tannins producing gallic acid which could be used for pharmaceutical applications. Bioremediation of heavy metals by employing biomass of A. glaucus will additionally help in tannery effluent treatment.

Conclusions

This study shows off the ability of tannase enzyme produced by a local fungal isolate, formerly identified as A. glaucus, in the bioremediation of tanneries containing heavy metals and tannin. Tannase enzyme was further subjected to partial purification by 80% ammonium sulfate saturation. The partial purified enzyme was characterized where found the optimum pH and temperature were 5 and 40 °C, respectively. Furthermore, the purified enzyme was active with more than 50% of its activity even at raised temperature as 60 °C which added a high-value for the enzyme usage in industrial bioprocesses require at high temperature. This can be considered as an additional advantage, since most of the processes assisted by tannase are performed at increased temperatures. Immobilized A. glaucus tannase was able to bio-convert tannic acid to gallic acid at 40 °C with tannic acid concentration up to 50 g/l. Using A. glaucus biomass at bioremediation of heavy metal from aqueous solution was effective where uptake percentages of (Cr3+, Pb2+, Cu2+, Fe3+, and Mn2+) were (37.20, 60.30, 55.27, 79.03 and 21.13% respectively). Additionally, the fungal biomass was able to repeatedly remove Cr3+ using desorbing agent (0.1 N HCl) for three cycles. The practical application of the metal biosorption process would be robustly improved when an efficient recovery of the sorbed metal and regenerated biomass is evolved for use in multiple sorption-desorption processes.

Data availability

The local isolated fungal strain was identified as Aspergillus glaucus and has a GeneBank accession number MH205732.

References

  1. Van de Lagemaat J, Pyle DL. Solid-state fermentation and bioremediation: development of a continuous process for the production of fungal tannase. Chem Eng J. 2001;84(2):115–23.

    Article  Google Scholar 

  2. Kumar M, Mehra R, Yogi R, Singh N, Salar RK, Saxena G, et al. A novel tannase from Klebsiella pneumoniae KP715242 reduces haze and improves the quality of fruit juice and beverages through detannification. Front Sustain Food Syst. 2023;7:1173611.

    Article  Google Scholar 

  3. Kumari M, Jain S. Tannins: an antinutrient with positive effect to Manage Diabetes. Res J Recent Sci [Internet]. 2012;1(12):70–3. Available from: www.isca.in.

    CAS  Google Scholar 

  4. Seth M, Chand S. Biosynthesis of tannase and hydrolysis of tannins to gallic acid by aspergillus awamori — optimisation of process parameters. Process Biochem. 2000;36(1–2):39–44.

    Article  CAS  Google Scholar 

  5. Mahendran B, Raman N, Kim DJ. Purification and characterization of tannase from Paecilomyces variotii: hydrolysis of tannic acid using immobilized tannase. Appl Microbiol Biotechnol [Internet]. 2006 Apr [cited 2024 Feb 16];70(4):444–50. https://pubmed.ncbi.nlm.nih.gov/16133325/.

  6. Monilola Wakil S, Damilola Ajayi O, Adedayo Fasiku S. Production of Tannase by Fungi isolated from different soils. J Basic Appl Sci. 2020;16:1–8.

    Article  Google Scholar 

  7. Papadaki E, Mantzouridou FT. Αpplication of Aspergillus niger for Extracellular Tannase and Gallic Acid Production in Non-sterile Table Olive Processing Wastewaters. Waste and Biomass Valorization [Internet]. 2023 Feb 1 [cited 2024 Feb 16];15(2):1199–212. https://link.springer.com/article/https://doi.org/10.1007/s12649-023-02242-0.

  8. Viswanath V, Leo VV, Prabha SS, Prabhakumari C, Potty VP, Jisha MS. Biosynthesis of tannase from cashew testa using Aspergillus niger MTCC5889 by solid state fermentation. J Food Sci Technol [Internet]. 2015 Nov 1 [cited 2023 Feb 2];52(11):7433–40. https://link.springer.com/article/https://doi.org/10.1007/s13197-015-1858-4.

  9. Roushdy MM, Desouky SE, Esmael ME, El-Louboudy SS, Elshikh HH. Optimization and Characterization of Tannin Acyle Hydrolase Produced by Aspergillus flavus var. columnaris Using Solid State Fermentation Technique. New York Sci J [Internet]. 2014 [cited 2024 Feb 16];7(3). http://www.sciencepub.net/newyork88:88-98].

  10. Reddy M, Reddy N, PRODUCTION OF TANNASE BY. ISOLATED ASPERGILLUS TERREUS UNDER SUBMERGED FERMENTATION. 2015.

  11. Roberta C, Juliana S, de Julyanna L, Jose CF, Jose EGG, de SSJ I, Keila AM, et al. Promising substrates to increase the production of tannase under solid state fermentation (SSF) by Penicillium spp. Afr J Biotechnol. 2017;16(45):2121–6.

    Article  Google Scholar 

  12. Chávez-González M, Rodríguez-Durán LV, Balagurusamy N, Prado-Barragán A, Rodríguez R, Contreras JC et al. Biotechnological Advances and Challenges of Tannase: An Overview. Food Bioprocess Technol [Internet]. 2012 Feb 27 [cited 2024 Feb 16];5(2):445–59. https://link.springer.com/article/10.1007/s11947-011-0608-5.

  13. Belmares R, Contreras-Esquivel JC, Rodríguez-Herrera R, Coronel AR, Aguilar CN. Microbial production of tannase: an enzyme with potential use in food industry. LWT - Food Sci Technol. 2004;37(8):857–64.

    Article  CAS  Google Scholar 

  14. Beniwal V, Chhokar V. Statistical Optimization of Culture Conditions for Tannase Production by Aspergillus awamori MTCC 9299 under Submerged Fermentation. Asian J Biotechnol [Internet]. 2009 Dec 15 [cited 2024 Feb 16];2(1):46–52. https://www.researchgate.net/publication/235624360_Statistical_Optimization_of_Culture_Conditions_for_Tannase_Production_by_Aspergillus_awamori_MTCC_9299_under_Submerged_Fermentation.

  15. Al-Mraai STY, Al-Fekaiki DF, Al-Manhel AJA. Purification and characterization of tannase from the local isolate of Aspergillus niger. J Appl Biol Biotechnol [Internet]. 2019 Jan 1 [cited 2024 Feb 16];7(1):29–34. https://www.researchgate.net/publication/329465876_Purification_and_characterization_of_tannase_from_the_local_isolate_of_Aspergillus_niger.

  16. Girdhari SN. Isolation and screening of tannase producing fungi. Int J Curr Microbiol Appl Sci. 2015;4(7):765–74.

    CAS  Google Scholar 

  17. Zhang YN, Yin JF, Chen JX, Wang F, Du QZ, Jiang YW et al. Improving the sweet aftertaste of green tea infusion with tannase. Food Chem [Internet]. 2016 Jul 17 [cited 2024 Feb 16];192:470–6. https://pubmed.ncbi.nlm.nih.gov/26304374/.

  18. Cavalcanti RMF, Jorge JA, Guimarães LHS. Characterization of Aspergillus fumigatus CAS-21 tannase with potential for propyl gallate synthesis and treatment of tannery effluent from leather industry. 3 Biotech [Internet]. 2018 Jun 1 [cited 2024 Feb 16];8(6):1–11. https://link.springer.com/article/10.1007/s13205-018-1294-z.

  19. Saxena G, Chandra R, Bharagava RN. Environmental pollution, toxicity profile and treatment approaches for tannery wastewater and its chemical pollutants. Rev Environ Contam Toxicol [Internet]. 2017 May 1 [cited 2024 Apr 30];240:31–69. https://link.springer.com/chapter/10.1007/398_2015_5009.

  20. Chaudhary P, Beniwal V, Kaur R, Kumar R, Kumar A, Chhokar V. Efficacy of Aspergillus fumigatus MCC 1175 for Bioremediation of Tannery Wastewater. CSAW [Internet]. 2019 Jun 1 [cited 2024 Apr 30];47(6):1900131. https://ui.adsabs.harvard.edu/abs/2019CSAW… C/abstract.

  21. Kumar R, Basu A, Bishayee B, Chatterjee RP, Behera M, Ang WL, et al. Management of tannery waste effluents towards the reclamation of clean water using an integrated membrane system: a state-of-the-art review. Environ Res. 2023;229:115881.

    Article  CAS  PubMed  Google Scholar 

  22. Gomes Júnior O, Santos MGB, Nossol ABS, Starling MCVM, Trovó AG. Decontamination and toxicity removal of an industrial effluent containing pesticides via multistage treatment: coagulation-flocculation-settling and photo-Fenton process. Process Saf Environ Prot. 2021;147:674–83.

    Article  Google Scholar 

  23. Duque AF, Campo R, Rio AV, Del, Amorim CL. Wastewater Valorization: Practice around the World at Pilot- and Full-Scale. Int J Environ Res Public Heal. 2021, Vol 18, Page 9466 [Internet]. 2021 Sep 8 [cited 2024 Feb 21];18(18):9466. https://www.mdpi.com/1660-4601/18/18/9466/htm.

  24. El-Gendy MMAA, El-Bondkly AMA. Evaluation and enhancement of heavy metals bioremediation in aqueous solutions by Nocardiopsis sp. MORSY1948, and Nocardia sp. MORSY2014. Brazilian J Microbiol. 2016;47(3):571–86.

    Article  CAS  Google Scholar 

  25. Khan AA, Mukherjee S, Mondal M, Boddu S, Subbaiah T, Halder G. Assessment of algal biomass towards removal of Cr(VI) from tannery effluent: a sustainable approach. Environ Sci Pollut Res [Internet]. 2021 Aug 25 [cited 2024 Feb 21];29(41):61856–69. https://link.springer.com/article/https://doi.org/10.1007/s11356-021-16102-8.

  26. Paria K, Pyne S, Chakraborty SK. Optimization of heavy metal (lead) remedial activities of fungi Aspergillus penicillioides (F12) through extra cellular polymeric substances. Chemosphere [Internet]. 2022 Jan 1 [cited 2024 Feb 21];286(Pt 3). https://pubmed.ncbi.nlm.nih.gov/34426280/.

  27. Ayele A, Haile S, Alemu D, Kamaraj M. Comparative Utilization of Dead and Live Fungal Biomass for the Removal of Heavy Metal: A Concise Review. Sci World J. 2021;2021.

  28. Gomaa OM, Jassim AY, Chanda A. Bioremoval of PVP-coated silver nanoparticles using Aspergillus niger: the role of exopolysaccharides. Environ Sci Pollut Res Int [Internet]. 2022 May 1 [cited 2024 Feb 21];29(21):31501–10. https://pubmed.ncbi.nlm.nih.gov/35001269/.

  29. Shalaby MA, Matter IA, Gharieb MM, Darwesh OM. Biosorption performance of the multi-metal tolerant fungus aspergillus sp. for removal of some metallic nanoparticles from aqueous solutions. Heliyon. 2023;9(5):e16125.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  30. Rose PK, Devi R. Heavy metal tolerance and adaptability assessment of indigenous filamentous fungi isolated from industrial wastewater and sludge samples. Beni-Suef Univ J Basic Appl Sci. 2018;7(4):688–94.

    Google Scholar 

  31. Saad AM, Saad MM, Ibrahim NA, El-Hadedy D, Ibrahim EI, El-Din AZK et al. Evaluation of Aspergillus tamarii NRC 3 biomass as a biosorbent for removal and recovery of heavy metals from contaminated aqueous solutions. Bull Natl Res Cent 2019 431 [Internet]. 2019 Jan 23 [cited 2024 Feb 16];43(1):1–9. https://bnrc.springeropen.com/articles/https://doi.org/10.1186/s42269-019-0046-5.

  32. Aftab K, Akhtar K, Jabbar A. Batch and column study for Pb-II remediation from industrial effluents using glutaraldehyde–alginate–fungi biocomposites. Ecol Eng. 2014;73:319–25.

    Article  Google Scholar 

  33. Karthick AS. Chemical Science Review and letters a review on methods, Application and properties of immobilized enzyme *Correspondence. Che Sci Rev Lett. 2012(3).

  34. Datta S, Christena LR, Rajaram YRS. Enzyme immobilization: an overview on techniques and support materials. 3 Biotech [Internet]. 2013 Feb [cited 2024 Feb 16];3(1):1. Available from: /pmc/articles/PMC3563746/.

  35. Hanachi P, Jafary F, Jafary F, Motamedi S. Immobilization of the Alkaline Phosphatase on Collagen Surface via Cross-Linking Method. Iran J Biotechnol [Internet]. 2015 Nov 1 [cited 2024 Feb 16];13(3):32–8. https://www.ijbiotech.com/article_11626.html.

  36. Chan LW, Lee HY, Heng PWS. Production of alginate microspheres by internal gelation using an emulsification method. Int J Pharm [Internet]. 2002 Aug 21 [cited 2024 Feb 16];242(1–2):259–62. https://pubmed.ncbi.nlm.nih.gov/12176259/.

  37. Wang L, Liu F, Jiang Y, Chai Z, Li P, Cheng Y et al. Synergistic antimicrobial activities of natural essential oils with chitosan films. J Agric Food Chem [Internet]. 2011 Dec 14 [cited 2024 Feb 16];59(23):12411–9. https://pubmed.ncbi.nlm.nih.gov/22034912/.

  38. Saad MM, Saad AM, Hassan HM, Ibrahim EI, Abdelraof M, Ali BA. Optimization of tannase production by Aspergillus glaucus in solid-state fermentation of black tea waste. Bioresour Bioprocess [Internet]. 2023 Dec 1 [cited 2024 Feb 16];10(1):1–14. https://bioresourcesbioprocessing.springeropen.com/articles/https://doi.org/10.1186/s40643-023-00686-9.

  39. la cerda gómez de. Valdés reyes, renTería meléndez, herrera rodríguez. Tannase producTion under solid and submerged culTure by xerophilic sTrains of aspergillus and their geneTic relaTionships. Micol Apl Int [Internet]. 2011;23(1):21–7. Available from: www.micaplint.com.

    Google Scholar 

  40. Wu C, Zhang F, Li L, Jiang Z, Ni H, Xiao A. Novel optimization strategy for tannase production through a modified solid-state fermentation system. Biotechnol Biofuels [Internet]. 2018 Apr 2 [cited 2024 Feb 16];11(1):1–15. https://biotechnologyforbiofuels.biomedcentral.com/articles/https://doi.org/10.1186/s13068-018-1093-0.

  41. MM B. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal Biochem [Internet]. 1976 May 7 [cited 2023 Feb 2];72(1–2):248–54. https://pubmed.ncbi.nlm.nih.gov/942051/.

  42. Özel YK, Gedikli S, Aytar P, Ünal A, Yamaç M, Çabuk A et al. Immobilization of cells and enzymes by gel entrapment. Immobil cells Enzym a Pract approach [Internet]. 1985 Oct [cited 2024 Feb 16];110(4):39–48. https://cir.nii.ac.jp/crid/1573105976030696832.

  43. Abdel-Naby MA, Sherif AA, El-Tanash AB, Mankarios AT. Immobilization of Aspergillus oryzae tannase and properties of the immobilized enzyme. J Appl Microbiol [Internet]. 1999 Jul 1 [cited 2024 Feb 16];87(1):108–14. https://doi.org/10.1046/j.1365-2672.1999.00799.x.

  44. SHILPA A. WAGH, BIOCONVERSION OF TANNIC ACID TO GALLIC ACID BY. USING FUNGAL TANNASE [Internet]. [Punne, India]: DIVISION OF BIOCHEMICAL SCIENCES; 2010 [cited 2023 Feb 2]. https://dspace.ncl.res.in/xmlui/bitstream/handle/20.500.12252/3759/TH1829.pdf?sequence=1&isAllowed=y

  45. Sharma S, Bhat TK, Dawra RK. A Spectrophotometric Method for Assay of Tannase using rhodanine. Anal Biochem. 2000;279(1):85–9.

    Article  CAS  PubMed  Google Scholar 

  46. Abd-Elmotey S, Nasar FR, Nasar SA, Abou-Taleb KA. Tannase and gallic acid production by Aspergillus niger SWP33 and Penicillium griseoroseum T11 using agricultural wastes under submerged and solid-state fermentation and its application. Egypt J Chem [Internet]. 2022 Sep 1 [cited 2023 Feb 2];65(9):21–35. https://ejchem.journals.ekb.eg/article_217540.html.

  47. Saad AM. Factors affecting cobalt uptake by cobalt-trained Mucor Rouxii NRRL 1894 biomass. Eur J Biotechnol Biosci. 2015;3(3):1–6.

    Google Scholar 

  48. Mahendran B, Raman N, Kim DJ. Purification and characterization of tannase from Paecilomyces variotii: hydrolysis of tannic acid using immobilized tannase. Appl Microbiol Biotechnol [Internet]. 2006 Apr [cited 2024 Apr 30];70(4):444–50. https://pubmed.ncbi.nlm.nih.gov/16133325/.

  49. Yao J, Guo GS, Ren GH, Liu YH. Production, characterization and applications of tannase. J Mol Catal B Enzym. 2014;101:137–47.

    Article  CAS  Google Scholar 

  50. Dhiman S, Mukherjee G, Singh AK. Recent trends and advancements in microbial tannase-catalyzed biotransformation of tannins: a review. Int Microbiol [Internet]. 2018 Dec 1 [cited 2024 Feb 18];21(4):175–95. https://pubmed.ncbi.nlm.nih.gov/30810902/.

  51. Abdulhameed Sabu GSK. and AP. Purification and Characterization of Tannin Acyl Hydrolase from Aspergillus niger ATCC 16620. Food Technol Biotechnol [Internet]. 2005 [cited 2023 Feb 2];43(2):582–8. https://www.researchgate.net/publication/228366317_Purification_and_Characterization_of_Tannin_Acyl_Hydrolase_from_Aspergillus_niger_ATCC_16620.

  52. Sabu A, Kiran GS, Pandey A. Purification and characterization of Tannin Acyl Hydrolase from Aspergillus Niger ATCC 16620.

  53. Govindarajan RK, Krishnamurthy M, Neelamegam R, Shyu DJH, Muthukalingan K, Nagarajan K. Purification, structural characterization and biotechnological potential of tannase enzyme produced by Enterobacter cloacae strain 41. Process Biochem. 2019;77:37–47.

    Article  Google Scholar 

  54. Kumar R, Sharma J, Singh R. Production of tannase from Aspergillus ruber under solid-state fermentation using jamun (Syzygium cumini) leaves. Microbiol Res [Internet]. 2006 Jul 25 [cited 2024 Feb 18];162(4):384–90. https://europepmc.org/article/med/16870410.

  55. Kasieczka-Burnecka M, Kuc K, Kalinowska H, Knap M, Turkiewicz M. Purification and characterization of two cold-adapted extracellular tannin acyl hydrolases from an Antarctic strain Verticillium sp. P9. Appl Microbiol Biotechnol [Internet]. 2007 Nov [cited 2024 Feb 18];77(1):77–89. https://pubmed.ncbi.nlm.nih.gov/17786433/.

  56. Wan Y, Fan H, Gao L, Li R, Xie M, Wu CF et al. The Change Mechanism of Structural Characterization and Thermodynamic Properties of Tannase from Aspergillus niger NL112 Under High Temperature. Appl Biochem Biotechnol [Internet]. 2021 Jul 1 [cited 2024 Apr 30];193(7):2225–44. https://pubmed.ncbi.nlm.nih.gov/33686629/.

  57. Mukherjee G, Banerjee R. Effects of temperature, pH and additives on the activity of tannase produced by a co-culture of Rhizopus oryzae and Aspergillus foetidus. World J Microbiol Biotechnol [Internet]. 2006 Mar 30 [cited 2023 Feb 2];22(3):207–12. https://link.springer.com/article/https://doi.org/10.1007/s11274-005-9022-3.

  58. Gayen S, Ghosh U. Purification and characterization of tannin acyl hydrolase produced by mixed solid state fermentation of wheat bran and marigold flower by penicillium notatum NCIM 923. Biomed Res Int. 2013;2013.

  59. Da Miura A, Kadowaki MK, Minozzo MC, Marques De Souza CG, Gandolfi Boer C, Bracht A et al. Production, purification and characterization of tannase from Aspergillus tamarii. African J Biotechnol [Internet]. 2012 Jan 5 [cited 2024 Feb 18];11(2):391–8. https://www.ajol.info/index.php/ajb/article/view/99229.

  60. Aharwar A, Parihar DK. Tannases: production, properties, applications. Biocatal Agric Biotechnol. 2018;15:322–34.

    Article  Google Scholar 

  61. Chhokar V, Seema, Beniwal V, Salar RK, Nehra KS, Kumar A et al. Purification and characterization of extracellular tannin acyl hydrolase from Aspergillus heteromorphus MTCC 8818. Biotechnol Bioprocess Eng [Internet]. 2010 Oct 9 [cited 2023 Feb 2];15(5):793–9. https://link.springer.com/article/https://doi.org/10.1007/s12257-010-0058-3.

  62. Keerti, Gupta A, Kumar V, Dubey A, Verma AK. Kinetic Characterization and Effect of Immobilized Thermostable β-Glucosidase in Alginate Gel Beads on Sugarcane Juice. ISRN Biochem [Internet]. 2014 Feb 20 [cited 2024 Feb 18];2014:1–8. https://pubmed.ncbi.nlm.nih.gov/25969764/.

  63. Aharwar A, Parihar DK. Talaromyces verruculosus tannase immobilization, characterization, and application in tea infusion treatment. Biomass Convers Biorefinery [Internet]. 2023 Jan 1 [cited 2024 Apr 30];13(1):261–72. https://link.springer.com/article/https://doi.org/10.1007/s13399-020-01162-6.

  64. Yu X, Li Y, Wang C, Wu D. Immobilization of Aspergillus niger tannase by microencapsulation and its kinetic characteristics. Biotechnol Appl Biochem [Internet]. 2004 Oct [cited 2024 Apr 30];40(Pt 2):151–5. https://pubmed.ncbi.nlm.nih.gov/14683525/.

  65. Sharma S, Agarwal L, Saxena RK. Purification, immobilization and characterization of tannase from Penicillium variable. Bioresour Technol [Internet]. 2008 May [cited 2024 Apr 30];99(7):2544–51. https://pubmed.ncbi.nlm.nih.gov/17570660/.

  66. Yao J, Chen Q, Zhong G, Cao W, Yu A, Liu Y. Immobilization and characterization of tannase from a metagenomic library and its use for removal of tannins from green tea infusion. J Microbiol Biotechnol [Internet]. 2014 [cited 2024 Apr 30];24(1):80–6. https://pubmed.ncbi.nlm.nih.gov/24150500/.

  67. Toldra F, Lequerica JL. Use of scanning electron microscopy techniques in the biocatalyst immobilization. Food / Nahrung [Internet]. 1991 Jan 1 [cited 2024 Apr 30];35(7):739–44. https://onlinelibrary.wiley.com/doi/full/10.1002/food.19910350717.

  68. Ye P, Xu ZK, Wu J, Innocent C, Seta P. Nanofibrous poly(acrylonitrile-co-maleic acid) membranes functionalized with gelatin and chitosan for lipase immobilization. Biomaterials [Internet]. 2006 Aug [cited 2024 Feb 18];27(22):4169–76. https://pubmed.ncbi.nlm.nih.gov/16584770/.

  69. Srivastava A, Kar R. Characterization And Application Of Tannase Produced By Aspergillus Niger ITCC 6514.07 On Pomegranate Rind. Braz J Microbiol [Internet]. 2009 [cited 2024 Apr 30];40(4):782–9. https://pubmed.ncbi.nlm.nih.gov/24031425/.

  70. Kanpiengjai A, Khanongnuch C, Lumyong S, Haltrich D, Nguyen TH, Kittibunchakul S. Co-production of gallic acid and a novel cell-associated tannase by a pigment-producing yeast, Sporidiobolus ruineniae A45.2. Microb Cell Fact [Internet]. 2020 Apr 25 [cited 2024 Feb 19];19(1):1–12. https://microbialcellfactories.biomedcentral.com/articles/https://doi.org/10.1186/s12934-020-01353-w.

  71. Srivastava S. K AD. Biological Wastes the Tool for Biosorption of Arsenic. J Bioremediation Biodegrad 2016 71 [Internet]. 2015 Dec 3 [cited 2024 Feb 19];7(1):1–3. https://www.omicsonline.org/open-access/biological-wastes-the-tool-for-biosorption-of-arsenic-2155-6199-1000323.php?aid=65328.

  72. Coelho LM, Rezende HC, Coelho LM, de Sousa PAR, Melo DFO, Coelho NMM et al. Bioremediation of Polluted Waters Using Microorganisms. Adv Bioremediation Wastewater Polluted Soil [Internet]. 2015 Sep 9 [cited 2024 Feb 19]; https://www.intechopen.com/chapters/48803.

  73. Kapoor A, Viraraghavan T, Cullimore DR. Removal of heavy metals using the fungus aspergillus Niger. Bioresour Technol. 1999;70(1):95–104.

    Article  CAS  Google Scholar 

  74. Dhankhar R, Hooda A. Fungal biosorption–an alternative to meet the challenges of heavy metal pollution in aqueous solutions. Environ Technol [Internet]. 2011 [cited 2024 Feb 20];32(5–6):467–91. https://pubmed.ncbi.nlm.nih.gov/21877528/.

  75. Gouda SA, Egypt. J Aquat Biol Fish. 2023;27(2):135–53.

    Article  Google Scholar 

  76. Saglam A, Yalcinkaya Y, Denizli A, Arica MY, Gene Ö, Bektas S. Biosorption of mercury by carboxymethylcellulose and immobilized Phanerochaete chrysosporium. Microchem J. 2002;71(1):73–81.

    Article  CAS  Google Scholar 

  77. Saad AM, Saad MM, Ibrahim NA, El-Hadedy D, Ibrahim EI, El-Din AZK et al. Evaluation of Aspergillus tamarii NRC 3 biomass as a biosorbent for removal and recovery of heavy metals from contaminated aqueous solutions. Bull Natl Res Cent 2019 431 [Internet]. 2019 Jan 23 [cited 2024 Apr 30];43(1):1–9. https://bnrc.springeropen.com/articles/https://doi.org/10.1186/s42269-019-0046-5.

  78. Singh R, Singh P, Sharma R. Microorganism as a tool of bioremediation technology for cleaning environment: a review. Proc Int Acad Ecol Environ Sci [Internet]. 2014;4(1):1–6. Available from: www.iaees.org.

    CAS  Google Scholar 

  79. Sakthi Priya N, Doble M, Sangwai JS. Bioremediation of Coastal and Marine Pollution due to Crude Oil using a Microorganism Bacillus subtilis. Procedia Eng. 2015;116(1):213–20.

    Article  Google Scholar 

  80. Ojuederie OB, Babalola OO. Microbial and Plant-Assisted Bioremediation of Heavy Metal Polluted Environments: A Review. Int J Environ Res Public Heal. 2017, Vol 14, Page 1504 [Internet]. 2017 Dec 4 [cited 2023 Feb 2];14(12):1504. https://www.mdpi.com/1660-4601/14/12/1504/htm.

  81. Sukumar C, Janaki V, Vijayaraghavan K, Kamala-Kannan S, Shanthi K. Removal of Cr(VI) using co-immobilized activated carbon and Bacillus subtilis: fixed-bed column study. Clean Technol Environ Policy [Internet]. 2017 Jan 1 [cited 2023 Feb 2];19(1):251–8. https://link.springer.com/article/10.1007/s10098-016-1203-2.

Download references

Acknowledgements

The authors would also like to acknowledge the facilities available at the National Research Centre, Cairo, Egypt, for providing the necessary research facilities.

Funding

Open access funding provided by The Science, Technology & Innovation Funding Authority (STDF) in cooperation with The Egyptian Knowledge Bank (EKB). Open access funding provided by The Science, Technology & Innovation Funding Authority (STDF) in cooperation with The Egyptian Knowledge Bank (EKB).

Open access funding provided by The Science, Technology & Innovation Funding Authority (STDF) in cooperation with The Egyptian Knowledge Bank (EKB).

Author information

Authors and Affiliations

  1. Microbial Chemistry Department, National Research Centre (NRC), 33 EL-Bohouth St., Dokki 12622, Giza, Egypt

    Moataza Mahmoud Saad, Abdelnaby Mahmoud Saad, Helmy Mohamed Hassan, Eman I. Ibrahim, Amany A. Hassabo & Basant A. Ali

Authors
  1. Moataza Mahmoud Saad
    View author publications

    You can also search for this author in PubMed Google Scholar

  2. Abdelnaby Mahmoud Saad
    View author publications

    You can also search for this author in PubMed Google Scholar

  3. Helmy Mohamed Hassan
    View author publications

    You can also search for this author in PubMed Google Scholar

  4. Eman I. Ibrahim
    View author publications

    You can also search for this author in PubMed Google Scholar

  5. Amany A. Hassabo
    View author publications

    You can also search for this author in PubMed Google Scholar

  6. Basant A. Ali
    View author publications

    You can also search for this author in PubMed Google Scholar

Contributions

All authors conceptualized and planned the research. Experiments were done, and data were evaluated by all authors. The manuscript was written by MMS, AMS, HMS, EII, AAH, and BAA. The article was reviewed by all authors. All authors read and approved the final manuscript.

Corresponding author

Correspondence to Basant A. Ali.

Ethics declarations

Ethics approval and consent to participate

There were no human or animal subjects used in any of the studies mentioned in this article.

Consent for publication

Not applicable.

Competing interests

The authors declare no competing interests.

Additional information

Publisher’s Note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Rights and permissions

Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by/4.0/. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated in a credit line to the data.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Saad, M.M., Saad, A.M., Hassan, H.M. et al. Bioremoval of tannins and heavy metals using immobilized tannase and biomass of Aspergillus glaucus. Microb Cell Fact 23, 209 (2024). https://doi.org/10.1186/s12934-024-02477-z

Download citation

  • Received:

  • Accepted:

  • Published:

  • DOI: https://doi.org/10.1186/s12934-024-02477-z

Keywords

Microbial Cell Factories

ISSN: 1475-2859

Contact us