Covalent Immobilization of Multi-Enzyme Biocatalysts on Nano-Silica for Efficient p,p'-DDT Degradation

doi:10.21203/rs.3.rs-3110783/v1

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Covalent Immobilization of Multi-Enzyme Biocatalysts on Nano-Silica for Efficient p,p'-DDT Degradation

https://doi.org/10.21203/rs.3.rs-3110783/v1

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Growing concerns over environmental pollution have necessitated the development of new-generation environmental protection technologies. Multi-enzyme biocatalysts offer a promising approach for reducing pollution caused by organic wastes. However, the recovery of free-form multienzymes is challenging, resulting in high costs and low production efficiency, limiting their application in bioremediation. Immobilization provides a solution by stabilizing and retaining enzymes. In this study, we devised a sensitive and stable enzyme biocatalyst by covalently immobilizing multi-enzymes onto nano-silica using glutaraldehyde. Ligninolytic enzymes (laccase, aryl alcohol oxidase, lignin peroxidase, and manganese peroxidase) were produced from Pleurotus ostreatus (NRRL-2366) under submerged fermentation. Enzymes were partially purified through ammonium sulfate precipitation and dialysis. These purified enzymes were immobilized on nano-silica. The resulting immobilized enzymes biocatalyst exhibited stability and activity across a pH range of 4 to 9 and a temperature range of 20 to 55°C. Immobilization of laccase, lignin peroxidase, manganese peroxidase, and aryl-alcohol oxidase achieved residual activities of 77%, 62.5%, 41.59%, and 28.21%, respectively, after three consecutive batches. Immobilized enzymes biocatalyst effectively degraded p,p'-DDT, and its complete degradation was achieved after incubation at pH 5 and 30°C for 12 hours, as confirmed by HPLC analysis. The GC-MS analysis revealed the detection of eleven major metabolites during the degradation process, which were utilized to predict the degradation pathway.

Multienzymes

Immobilization

Biocatalyst

p'-DDT

Biodegradation

Silica nanoparticles

Water is a vital resource that supports all life on Earth. However, concerns about the shortage of freshwater supply and its impact on the sustainability of human societies have been increasing in recent decades (Rathi et al. 2021). In particular, there has been growing attention to the occurrence and significance of persistent organic pollutants (POPs) in marine environments. POPs are widely distributed, persistent, and toxic compounds that pose various health risks to both humans and the environment. Exposure to these pollutants can result in a wide range of adverse health effects, ranging from acute symptoms such as allergies, hypersensitivity, and dermatological reactions, to more severe consequences like endocrine disruption, reproductive and immune dysfunction, neurological disorders, and cancer. Organochlorine pesticides, including DDT (1,1,1-trichloro-2,2-bis(4-chlorophenyl) ethane), have been extensively used in agriculture and public health since the 1940s (Purnomo et al. 2017). DDT was initially introduced as an organochlorine pesticide during World War II to combat disease-spreading mosquitoes, gaining global popularity as the first synthetic pesticide. Although banned in most countries due to its negative impacts on wildlife and human health(Auteri et al. 2022), DDT is still used for essential public health and agricultural purposes in some developing nations (Foght et al., 2001). Despite the ban, DDT residues remain present in the environment, particularly in flowing water bodies and agricultural soil (Zdarta et al. 2018). Physicochemical processes are commonly employed for DDT removal; however, these methods often involve harsh reaction conditions, generate free radicals and secondary pollutants, and are associated with high operational costs (Zhang et al. 2013). Therefore, bioremediation processes offer an attractive alternative. Bioremediation strategies are cost-effective and enable the transformation of DDT and its major byproducts into harmless substances through mineralization (Ahmed et al. 2001).

Due to its eco-friendliness and biodegradability, enzymatic degradation of DDT is a favorable approach. However, the free enzyme encounters several limitations, such as poor stability, lack of reusability, and high cost, which hinder its practical application. Immobilization of enzymes, where they are attached to solid supports, can address these limitations by preserving the enzyme's structure and minimizing product contamination (Zdarta et al. 2018; Kiran et al. 2020). Immobilized enzymes offer advantages such as easy recovery, low energy consumption, cost-effectiveness, operational stability, and reusability (Zdarta et al. 2018; Habimana et al. 2021). Multi-enzyme biocatalysis is a significant technology used in various industries to produce valuable chemicals. Immobilization of multi-enzymes on support materials has proven to be an efficient approach, enhancing enzymatic activity through substrate channeling and improving enzyme stability and reusability. Various materials, including graphene, carbon nanotubes, metal-organic frameworks, DNA nanostructures, polymers, and silica, have been employed for multi-enzyme immobilization, providing protection against heavy metals, high temperatures, and challenging biological conditions (Ren et al. 2019; Alcántara et al. 2022).

Silica nanoparticles have emerged as promising candidates for immobilization due to their high surface area, pore volume, and lack of UV absorption. Silica and silicon nanomaterials find applications in drug/gene delivery, lightweight aggregates, regenerative medicine, tissue engineering, cancer diagnosis/therapy, catalysis, and energy storage. The demand for eco-friendly synthesis of silica and silicon nanomaterials has increased, focusing on cost-effectiveness, safety, reliability, simplicity, scalability, and controllable particle size distribution. These materials can be obtained from sustainable agricultural resources like rice husk ash, sugar cane, coffee husk, wheat husk, and corn cob ash. The use of green and sustainable techniques eliminates or reduces complex procedures and minimizes environmental contaminants, facilitating the production of advanced materials for high-tech applications (Alcántara et al. 2022).

The utilization of nanosilica and multi-enzymes offers a promising solution to overcome the limitations associated with free-form enzymes. Nanosilica provides a stable and efficient support material, while immobilization enhances enzyme stability and activity. This approach addresses the challenges of enzyme recovery, high costs, and low production efficiency, making it a valuable tool for bioremediation and environmental protection.

2.1. Chemicals and media

The standard samples of p,p'-DDT (purity 99.5%), p,p'-DDD (purity 98.0%), p,p'-DDE (purity 98.5%), and DDA (purity 99.0%) were acquired from the Central Agricultural Pesticide Laboratory (CAPL), Agricultural Research Center, Egypt. Analytical-grade sodium sulfate anhydrous, ethyl acetate, n-hexane, and glutaraldehyde were obtained from the Aldrich Company.

2.2.1. Fungal strain and culture conditions

The white-rot fungus Pleurotus ostreatus (NRRL-2366) was obtained from the National Central of Agricultural Utilization Research Service, U.S. Department of Agriculture, located in Peoria, Illinois, USA. The fungus was stored at a temperature of 4 ± 1°C on potato dextrose agar (PDA) slants and sub-cultured every 30 days on fresh medium. Subsequently, it was incubated at 25 ± 1°C for a period of 7–8 days to facilitate growth.

The cultivation medium for the production of ligninolytic enzymes was prepared following the method described by (Alcántara et al. 2022). Pleurotus ostreatus NRRL-2366 was grown in a 3-liter medium with the following composition (g/l): (NH₄)₂SO₄ (5.0), yeast extract (5.0), KH₂PO₄ (2.0), and MgSO₄.7H2O (0.3). The medium was supplemented with rice straw (10 g/l) as a carbon source instead of glucose. The cultivation was carried out for 14 days at 28°C and 200 rpm. To inoculate the medium, mycelium from 8-day-old cultures in 1–1 Roux flasks, containing 100 ml of the same medium, was washed with water, homogenized, and used as inoculum at a concentration of 1 g/l.

2.2.2. Purification of ligninolytic enzymes

The purification process of ligninolytic enzymes involved several steps. Firstly, extrudates were obtained by squeezing fermented rice straw through muslin cloth and then centrifuged at 8000 rpm and 4°C for 15 minutes. To obtain crude enzymes, 3 liters of the culture liquid from a 14-day-old culture were concentrated six-fold using ultrafiltration with a 10-kDa cut-off membrane. Polysaccharides were removed from the concentrated solution as described in previous studies (Guillén et al. 1990). The resulting supernatant was then subjected to ammonium sulfate saturation to precipitate the proteins. After adding an ammonium sulfate solution and incubating at 4°C for 30 minutes, the precipitates were collected by centrifugation at 10,000g for 10 minutes at 4°C. The precipitates were subsequently dissolved in 2.5 ml of sodium acetate buffer (25 mM, pH 5.5), and the protein content and enzyme activity were measured at each step. The active fractions obtained from precipitation at 30–80% ammonium sulfate saturation were combined and subjected to dialysis purification using a dialysis membrane with a molecular weight cut-off of 10,000 Dalton at 4°C in a 25 mM sodium acetate buffer (pH 5.5). The dialysate samples were used to determine protein and ligninolytic enzyme activities (Gaonkar and Furtado, 2020).

2.2.3. Enzymatic activities and protein content

The enzymatic activities and protein content were measured using culture supernatants at room temperature. The results represent the average of three independent assays. Enzymatic activity was expressed in International units (U), which represents the amount of enzyme needed to produce 1 µmol of product per minute and is expressed as U/L.

Laccase (Lac) activity was determined following the method described by (Johannes and Majcherczyk 2000). The substrate used was 2,2'-azinobis-3-ethylbenzthiazoline-6-sulfonate (ABTS), and the oxidation of ABTS was measured by the increase in absorbance at 420 nm. The extinction coefficient used (λ mM = 36 mM^− 1cm^− 1). One unit of enzyme activity was defined as the amount of enzyme required to oxidize 1 mmol of ABTS per minute, and the reactions were conducted at 30°C.

Aryl-alcohol oxidase (AAO) activity was determined spectrophotometrically by measuring the oxidation of veratryl alcohol to veratraldehyde. The reaction mixture consisted of 0.1 M sodium phosphate buffer (pH 6.0) and 10 mM veratryl alcohol (Okamoto and Yanase 2002). Lignin peroxidase (LiP) activity was measured following the method of Tien and Kirk (1984), which involved the H₂O₂-dependent oxidation of veratryl alcohol to veratraldehyde at 25°C and 310 nm. Manganese peroxidase (MnP) activity was measured using 2,6-dimethoxyphenol at 469 nm and 25°C, as described by Kuwahara et al. (1984).

For all enzyme activities, 1 U of enzyme activity was defined as the amount of enzyme releasing 1 µmol of oxidized product per minute at 25°C. The total protein content was determined using the Bradford method (1976) with bovine serum albumin as the standard.

2.3.1. Synthesis of nano silica powder from rice husk:

The synthesis of nano-silica powder from rice husk was carried out following the methodology described by (Rafiee et al. 2012). Initially, rice husk (RH) was washed with deionized water and dried in an oven at 105 ºC to remove moisture. The dried RH was then calcinated at 700 ºC for 6 hours to obtain silica. Ten grams of the calcinated RH were mixed with 80 mL of a 3.0 M NaOH solution and boiled for 3 hours. The resulting solution was filtered, and the residue was washed with 20 mL of deionized water. The filtrate was allowed to cool to room temperature, and 2.5 M H₂SO₄ acid was added until the pH of the solution reached 2. Subsequently, NH₄OH was added until the pH reached 8.5 at room temperature. The filtrate was dried at 120 ºC for 12 hours, and the resulting sample was powdered. Pure silica was extracted from the sample by refluxing with 6 M HCl for 4 hours, followed by repeated washing with deionized water to remove acidity. The purified silica was separated by centrifugation and dried at 105 ºC for 2 hours. The pure silica sample obtained from RH was then dissolved in 3.0 M NaOH through continuous stirring for 10 hours. The pH was adjusted to the range of 7.5–8.5 by adding 0.5 M H₂SO₄ drop-wise. The precipitated silica was washed repeatedly with warm deionized water until the filtrate was free from alkali. The precipitate was separated by centrifugation (9000 rpm for 1 hour), and the resulting sample was dried at 50 ºC for 48 hours to obtain nano-silica. Finally, the prepared nano-silica sample was used for immobilizing different catalyst enzymes.

2. 3.2. Covalent immobilization of oxidoreductase enzymes on nano-silica powder:

In the immobilization process, 0.5g of rice husk silica (RHS) nanocomposite was subjected to multiple washes with 50 mM sodium phosphate buffer (pH 7.2). Subsequently, a solution of 1% glutaraldehyde (GDA) in 50 mM sodium phosphate buffer (pH 7.2) was added to 2.0 g of nano-silica, and the mixture was gently shaken at 25°C for 4 hours to create GDA-linked nano-silica. The proteins were then covalently immobilized onto the GDA-linked nano-silica by incubating them together at 25°C for 24 hours with gentle shaking. Afterward, centrifugation at 3000 g was performed to recover the immobilized proteins from the solution. To remove nonspecifically adsorbed proteins, the matrix was washed with 1 M NaCl and 1% Tween 20, followed by 50 mM sodium phosphate buffer (pH 7.2) (Le et al., 2018). The mass of the immobilized proteins was determined by subtracting the mass of the non-immobilized proteins from the total mass. The immobilization efficiency was calculated as the percentage of immobilized protein relative to the total protein.

2.3.3 Characterization of synthesized materials.

The synthesized Nano-SiO₂ and Nano-SiO₂-Enz materials were characterized using scanning electron microscopy (SEM) and Fourier transform infrared spectroscopy (FTIR). For SEM analysis, the materials' surface morphology was examined using a Jeol JSM-6360LA SEM at a low magnification. The synthesized powder samples were fixed onto a specimen holder and coated with a layer of gold using the sputtering method. SEM images were captured at a magnification of 1000× with a 10 kV accelerating voltage.

To analyze the functional groups present on the surfaces of Nano-SiO2 and Nano-SiO2-Enz, Fourier transform infrared spectroscopy (FTIR) was performed. The data were obtained using a Bruker Vertex 80 spectrometer equipped with a Ram-FT module (RAM II). Diffused reflectance style was employed for data generation during FTIR analysis.

2.3.4. Effect of pH and temperature on the activity and stability of thr immobilized biocatalyst

The influence of pH on the activities of immobilized enzymes (Lac, AAO, LiP, and MnP) was assessed using spectrophotometric methods. The activities of the immobilized enzymes were measured in various buffer solutions, including 0.05 M Glycine-HCl buffer (pH 3, 5, 7, 8), 0.05 M Sodium acetate buffer (pH 4, 5, 6), and 0.05 M Tris-base buffer (pH 8, 9). To determine the activity of immobilized enzymes, 30 mg of the immobilized enzyme was used (Almeida et al. 2018).

The stability of nano-silica immobilized enzymes at different pH levels was also investigated following a similar method as described by (Almeida et al. 2018). Both free and immobilized enzymes were incubated at 25°C in various buffers, including 0.2 M sodium acetate buffers (pH 4.0, 5.0, and 6.0) and 0.2 M phosphate buffer (pH 7.0, 8.0, 9.0, and 10.0), for 3 hours. The residual activities of the enzymes were then determined.

Furthermore, the effect of temperature on enzyme activities was examined in the range of 20–70°C. Both free enzymes and immobilized enzymes were incubated for 20 minutes at temperatures ranging from 20 to 70°C in a buffer with pH 7.0. The residual activities were evaluated at 25°C after cooling. The results for pH and temperature were normalized, with the highest value in each set being assigned 100% activity (Liu et al. 2012).

2.3.5. Reusability assessment of the immobilized biocatalyst

To assess the reusability of immobilized enzymes obtained using different methods, 0.2 g of the immobilized enzyme was added to an enzyme substrate adjusted to the appropriate pH and incubated at the optimal temperature. Following the initial reaction, the immobilized enzyme was washed with Na-citrate buffer at pH 7.0, and its activity was measured. This process was repeated, and the activity was measured after each subsequent reaction cycle (Rahim et al. 2013). Throughout all stability experiments, the residual enzymatic activity was determined using the aforementioned method, and the initial activity of all enzymes was considered as 100%. This allowed for the comparison of immobilized enzyme activity over multiple cycles and the assessment of their reusability.

2.4. Biodegradation performance of p,p'- DDT using immobilized of oxidoredctase multi- enzymes

2.4.1. Optimal reaction pH conditions:

To determine the optimal reaction pH conditions, test tubes were prepared with varying pH buffers (5 in total). Each test tube contained 0.5 g of immobilized enzyme and 10.0 mg/L of p,p'-DDT. The test tubes were then incubated in a 30°C water bath for a period of 12 hours. The purpose of this experiment was to evaluate the efficiency of the immobilized oxidoreductase multi-enzymes in degrading p,p'-DDT at different pH levels.

2.4.2. Analysis and measurements

The concentration of p,p'-DDT in the reaction mixture was determined using high-performance liquid chromatography (HPLC) with a UV-Visible Diode Array detector (CECIL 4100, USA). To separate the compounds, a Nucleodur Sphinx RP column (25.0 cm × 4.6 mm, MZ-1 PerfectSil, Germany) was employed at a flow rate of 1 ml/min. A gradient elution method was applied by utilizing a mixture of acetonitrile and 0.5% acetic acid (50:50, v/v) over a period of 10 minutes. All experiments were conducted in triplicate to ensure the accuracy and reliability of the obtained results.

2.4.3 Extraction and determination of metabolites

After a 6-hour incubation period, the metabolites of p,p'-DDT were extracted from the supernatant of the cultured cell suspensions that were incubated with p,p'-DDT and p,p'-DDE in a reaction mixture at a concentration of 10 mg/l. To prepare the metabolites for identification, 5 ml of the reaction mixture supernatant was initially extracted with 5 ml of hexane, followed by an additional extraction with 5 ml of ethyl acetate. The organic phase was collected, dehydrated using anhydrous sodium sulfate, and the extracts were combined. After drying with a nitrogen blowing instrument, the volume was adjusted to 5 ml with hexane (Xie et al. 2018).

The metabolites were analyzed using a Clarus 500 GC/MS system (PerkinElmer, USA) equipped with an autosampler, a splitless model injector, and an Elite-5MS capillary column (crosslinked 5% phenyl-95% methyl silicone, 30 m × 0.25 mm × 0.25 µm). The initial oven temperature was set to 70°C with a hold time of 1.0 minute, followed by an increase to 180°C at a rate of 20°C/min and a hold time of 5.0 minutes. The temperature was then increased by 5°C per minute until reaching 260°C. The injector temperatures were maintained at 250°C. Electron ionization with an electron energy of 70 eV was employed, and the ion source temperature of the mass spectrometer was set to 250°C. The scanning range for the mass spectrometer was from 40 to 550 µ. The transfer line temperature was maintained at 280°C. The metabolites were identified by comparing their retention times and mass spectra with known reference samples. This comparison involved utilizing the mass spectral data provided in the library of WILEY, National Institute of Standard and Technology (NIST-11), as well as the mass spectral data available for p,p'-DDT metabolites in the literature. By examining the retention times and mass spectra, the metabolites were successfully detected and identified.

3.1. Ligninolytic enzymes production:

Rice straw (1%) was used as a carbon source for fermentation trials. Submerged fermentation of Pleurotus ostreatus NRRL-2366 was conducted at a temperature of 28ºC for a duration of 14 days. Figure 1 illustrates the ligninolytic enzyme production ability of the screened strain Pleurotus ostreatus NRRL-2366, including Lac, AAO, LiP, and MnP. The highest activities for Lac, AAO, LiP, and MnP were recorded on the 14th day, with values of 56.60 ± 2.83 U/ml, 13.48 ± 0.55 U/ml, 10.96 ± 0.55 U/ml, and 11.09 ± 0.55 U/ml, respectively. White-rot fungi (WRF) possess a unique extracellular ligninolytic system that oxidizes and breaks down lignin, and P. ostreatus exhibits the ability to produce both hydrolytic and ligninolytic enzymes simultaneously during the fermentation of ligninocellulosic materials (Lundell et al. 2010; Ozcirak Ergun and Ozturk Urek 2017)

Table 1 Partial purification of Ligninolytic enzymes from Pleuratus osteratus.

Ligninolytic enzymes	Purification steps	Total volume (mL)	enzyme activity (U)	Protein (mg)	Specific activity (U/mg)	Purification fold	% Yield
Lac	Crude Extract	500	28300	166.22	170.481	1.0	100
Lac	Ammonium sulphate precipitation	50	1750	6.86	255	1.5	6.18
AAO	Crude Extract	500	6740	166.22	40.548	1.0	100
AAO	Ammonium sulphate precipitation	50	407	6.86	59.324	1.46	6.03 60.38
LiP	Crude Extract	500	5480	166.22	32. 968	1.0	100
LiP	Ammonium sulphate precipitation	50	321	6.86	46.79	1.4	5.86
MnP	Crude Extract	500	5545	166.22	33.359	1.0	100
MnP	Ammonium sulphate precipitation	50	378	6.86	55.102	1.7	6.81

- Each value represents the mean ± S.D (Standard Division) and mean of three replicates

- Values in the same column with the same letter are not significantly at a significance level of P ≤ 0.05

- Lac represents Laccase, AAO represents Aryl-alcohol oxidase, LiP represents Lignin peroxidase, and MnP represents Manganese peroxidase.

3.1. Characterization of the synthesized nano-silica

SEM images were taken to examine the nanoparticles with and without immobilized catalyst enzyme (Fig. 2a, 2b). Figure 2a shows that the pure SiO₂ nanoparticles were uniform, loose, and without aggregation, with a grain size ranging from 5nm to 100nm. In contrast, the SEM images of the nano-SiO₂ catalyst enzyme (Fig. 2b) revealed numerous small crystal materials tightly adhered to the surface, with irregular crystal arrangement and micro holes present. This observation suggests that the SiO₂ particles adsorbed a significant amount of catalyst enzymes, resulting in their irregular appearance. The nano-silica particles appeared agglomerated and in an amorphous form without clear boundaries. Hence, the SEM analysis confirmed that the size of the silica particles synthesized from rice husk falls within the nano range of 1-100 nm.

The FTIR spectrum of the nano silica powder and the nano-silica immobilized with catalyst enzyme is shown in Fig. 3(a, b). Figure 3(a) exhibits strong absorption peaks at 463, 621, and 791 cm^− 1, corresponding to the Si–O–Si bending, Si–H, and symmetric Si–O–Si stretching modes of vibrations, respectively. These peaks confirm the presence of highly pure nano silica powder obtained from rice husk using a 2.5N NaOH purification treatment. Additionally, a peak at 1069 cm^− 1 is observed, which is assigned to Si-O-Si asymmetric stretching vibration. This further supports the amorphous nature of the nano-silica synthesized from rice husk. In contrast, Fig. 3(b) displays additional functional groups at 1348, 1124, 992, and 888 cm^− 1, which are attributed to sulfonamide groups (S = O), secondary alcohol (C-O), alkene (C = C), and alkene (C = C) groups, respectively. These additional functional groups indicate the presence of the immobilized catalyst enzymes.

The findings suggest that the enzymes laccase (Lac), aryl alcohol oxidase (AAO), lignin peroxidase (LiP), and manganese peroxidase (MnP) derived from Pleurotus ostreatus (NRRL-2366) were successfully immobilized onto nano-silica supports through the utilization of glutaraldehyde as a cost-effective and straightforward method. This immobilization technique, which involves activating the supports with amino groups and employing a glutaraldehyde spacer arm, is commonly used in the biotechnology field to enhance the stability of various enzymes for diverse applications (Hwang et al. 2004; Barbosa et al. 2013). The overall process is depicted in Fig. 4.

The efficiency of enzyme immobilization was assessed by determining the immobilization-activity yields, which were found to be 86.22 ± 4.55%, 76.11 ± 3.22%, 78.16 ± 1.82%, and 60.21 ± 3.98% for laccase (Lac), aryl alcohol oxidase (AAO), lignin peroxidase (LiP), and manganese peroxidase (MnP), respectively (Fig. 5). Covalent binding, using glutaraldehyde as a spacer, offers several advantages in terms of enzyme stability and minimized enzyme leaching. The ordered mesoporous silica provides an ideal platform for covalent immobilization due to its well-defined silanol groups, which offer reactive sites for enzyme functionalization and adjustable surface properties. This enables precise control over the positioning and density of the immobilized catalyst (Alptekin et al. 2011; Nadar et al. 2016a; Abdel-Sater et al. 2019). Furthermore, the five-atom carbon chain of glutaraldehyde serves as a spacer for enzymes, facilitating easier accessibility of their active sites to substrates. Glutaraldehyde exhibits rapid reactivity with amine groups, particularly at neutral pH (7), making it more effective than other aldehydes in forming crosslinks that are both thermally and chemically stable (Nimni et al. 1987).

3.2. Biochemical Characterization of nano-silica immobilized enzymes biocatalyst

The impact of pH on the activities of the immobilized enzymes (Lac, AAO, LiP, and MnP) was examined using spectrophotometric analysis. The activities of the immobilized enzymes were measured in various buffer solutions, including 0.05 M Glycine-HCl buffer (pH 3, 5, 7, 8), 0.05 M Sodium acetate buffer (pH 4, 5, 6), and 0.05 M Tris-base buffer (pH 8, 9). A total of 30 mg of immobilized enzyme was used for the activity assays. To evaluate the stability of the immobilized biocatalyst at different pH levels, the immobilized enzymes were incubated in buffer solutions ranging from pH 3.0 to 10.0 for 3 hours at room temperature. After incubation, the residual activity of each enzyme was determined using the standard assay method. As depicted in Fig. 6a, purified laccase (1 mg/mL) exhibited maximum activity at pH 3.0, whereas the immobilized enzyme (30 mg/mL) showed a shift of one unit towards higher pH values after cross-linking of laccase on nanosilica. This shift can be attributed to the alteration of the microenvironment of laccase by nanosilica, including ionic interactions between the enzyme and charged surfaces of nanosilica. According to (Jiang et al. 2005), immobilization can create a microenvironment around the enzyme where unequal partitioning of H⁺ and OH⁻ concentrations, resulting from electrostatic interactions with the support, can cause

A change in the optimal pH was observed after covalent immobilization of laccase on different supports. For instance, a shift of 0.5 and 1.5 units towards higher pH values was reported when laccase was immobilized on Fe₃O₄-CS-CCn and Fe₃O₄-CS-EDAC supports, respectively (Kalkan et al. 2012). (B. Yang et al. 2021) also demonstrated a shift in the optimum pH for the oxidation of catechol upon immobilization of laccase on magnetic mesoporous nanosilica. In the pH stability test, it was found that the relative activity of the immobilized laccase was retained at approximately 30% at pH 8.0, while the purified laccase was inactive. The silica-laccase nanoparticles exhibited over 90% residual activity within the pH range of 3–5, whereas the purified laccase showed a significant decrease in residual activity within the same pH range (Fig. 7a). Similar findings have been reported by several researchers, indicating that the pH stability of laccase improved within the pH range of 2.0–7.0 after immobilization on DEAE-Granocel 500 and sol–gel matrix supports (Kalkan et al. 2012; Muhammad Nasir Iqbal and Asgher 2013; Gahlout et al. 2017).

The catalytic rates of immobilized aryl alcohol oxidase (AAO) on nano-silica (30 mg) were compared to those of free AAO (1 mg/mL) at various pH values under the same conditions (Fig. 6b). The optimal activity for the immobilized enzyme was observed at pH 6.0–7.0, with relatively high activities retained up to pH 8.0. In contrast, the free enzyme exhibited the highest reaction rate at pH 6.0, and a 50% reduction in activity was observed at pH 8.0. Under acidic conditions (pH 3–4), both free and immobilized AAO enzymes showed similar low activities. These findings suggest that the immobilized AAO enzyme exhibited improved pH stability compared to the free enzyme under alkaline conditions. The pH stability test revealed that almost 35% of the relative activity of immobilized AAO was retained at pH 3, while the purified AAO was inactive. Furthermore, the AAO nanoparticle exhibited over 90% residual activity in the pH range of 6–8, whereas the purified AAO showed a comparable decrease in residual activity within the same pH range (Fig. 5b). These results are consistent with the findings of Kato et al. 2009, who reported improved pH stability of AAO immobilized on 3D-mesoporous silicate materials compared to the free enzyme. AAO is primarily active within a neutral to slightly acidic pH range (5.5–7.5), with only a few enzymes active in more alkaline conditions (Goswami et al. 2013). Immobilization of AAO offers new possibilities for its catalytic activity and stability, opening doors for industrial and biocatalytic applications (Jun Kim et al. 2001; Bühler and Schmid 2004; Liu et al. 2021).

Figure 6c illustrates the effect of pH conditions on the activity of free lipase (Lip) at 1 mg/mL and immobilized Lip at 30 mg. The results showed that free Lip had maximum activity at pH 3–5, while the highest catalytic activity of nano-silica immobilized Lip ranged between pH 5–7. Moreover, the immobilized Lip exhibited strong acid-base tolerance (Fig. 6c). The relative activity of immobilized Lip remained above 90.0% in the pH range of 6–7 and reached 80% and 74% in acidic (pH 4.0) and basic (pH 8.0) solutions, respectively. This enhanced pH stability can be attributed to the protection provided by the carrier and the altered protein structure of the immobilized enzyme, which reduces the influence of pH factors (Kiani et al. 2022).

Free manganese peroxidase (MnP) remained relatively stable in the pH range of 4–6, but rapid inactivation occurred outside of this range (Fig. 6d). Previous studies have reported that LiPs from white rot fungi typically exhibit higher activities between pH 2 and 5, and after immobilization, a slight shift towards more acidic conditions is observed (Asgher et al. 2012). However, LiP from Ganoderma lucidum (GRM117) exhibits optimal pH in basic conditions after immobilization (Oliveira et al. 2018). The purified MnP also remained comparatively stable in the pH range of 4.0–6.0 but was rapidly inactivated outside of this range. The enzyme is susceptible to inactivation at pH 6.5 and higher

The temperature effects on both free and immobilized enzymes were investigated in the range of 20–70°C under standard conditions for each enzyme. The optimum temperature for all free and immobilized enzymes was found to be within the range of 25–30°C (Figs. 7a, b, c, and d). After cross-linking laccase on nanosilica, there was an observed shift in the optimum temperature towards higher values. The purified laccase had an optimum temperature of 25°C, while the immobilized laccase showed optimum activity at 30°C.

To assess the thermal stability of the enzymes, the residual activity was measured after incubating the free and the immobilized biocatalyst in the absence of substrate at different temperatures (20–70°C) for 20 minutes (Figs. 7a and b). The immobilized biocatalyst demonstrated superior stability, retaining 92%, 83%, 76%, and 63% residual activity after incubation at 55°C for laccase, AAO, lipase, and MnP, respectively. However, the immobilized biocatalyst became rapidly inactivated at temperatures above this range. In comparison, the free enzymes retained only 80%, 58%, 45%, and 38% residual activity at the same temperature. The enhanced thermo-stability of the immobilized biocatalyst can be attributed to the glutaraldehyde-enzyme cross-linking, which prevents conformational distortion of the enzyme at high temperatures (Sojitra et al. 2016; Panwar et al. 2017).

In a simplified view, this strategy achieves multiple intramolecular cross-linking, where the cross-linking agent is a flat and multifunctional structure formed by the activated support surface. This approach enhances enzyme stability against various destabilizing factors, including heat, solvents, and chaotropic reagents (Inoue et al. 2019; Parui and Jana 2021; C. Yang et al. 2021). The potential of multipoint covalent immobilization to stabilize enzymes is widely accepted in the scientific community (Weltz et al. 2020). This strategy relies on maintaining the fixed positions of all groups involved in immobilization under any experimental condition, allowing only limited movement determined by the length of the spacer arm. This contributes to improved enzyme stability by reducing conformational changes. Furthermore, it is worth mentioning that cold denaturation, a distinct phenomenon from thermoinactivation, can also induce conformational changes leading to inactive forms of enzymes (C. Yang et al. 2021).

3.3. Reusability assessment of the immobilized enzymes biocatalyst

The reusability of the immobilized biocatalyst was evaluated, as depicted in Fig. 7. It was found that the immobilized Lac maintained approximately 77% of its activity after the third cycle, while AAO, Lip, and MnP retained approximately 62.5%, 41.59%, and 28.21% of their activities, respectively. The biocatalyst could be reused for up to five consecutive cycles at a temperature of 30°C.

One of the primary objectives of enzyme immobilization is to enable multiple reuse cycles, thereby reducing costs in industrial processes (Abdel-Sater et al. 2019). The ability to reuse the immobilized enzyme for multiple cycles may be attributed to factors such as multipoint, multi-subunit immobilization or the creation of favorable microenvironments (Xu et al. 2014). However, a decrease in activity over successive cycles could be attributed to denaturation, enzyme leakage during use, and diffusional effects (Ye et al. 2006).

3.4.1. p,p'- DDT degradation efficiency by the immobilized enzymes biocatalyst:

To assess the efficiency of p,p'-DDT degradation, the optimal conditions determined from previous experiments were employed. The remaining residual amounts of p,p'-DDT were quantified using HPLC analysis.

Figure 8 illustrates that the immobilized multi-enzyme biocatalyst achieved complete degradation of p,p'-DDT (10 mg L-1) under the optimized conditions of pH 5.0, incubated at 30°C for 24 hours.

To validate the degradation of p,p'-DDT by the immobilized enzymes biocatalyst and gain insights into potential degradation pathways, the extracted samples were subjected to GC-MS analysis. The results are summarized in Table 2, and the corresponding figures (Figs. 10 and 11) provide visual representations of the findings.

The GC-MS analysis revealed the presence of eleven major metabolites in the experimental samples, including actual and predicted degradation products. These metabolites were compared to the parent p,p'-DDT, which had a molecular weight (m/z) of 354 and a retention time (RT) of 36.98 min. The authentic standard of p,p'-DDT listed in the library databases and the control pure compound showed identical characteristics. Notably, the mass-spectral analysis of the samples indicated the presence of a metabolite with a parent ion of m/z 316 and a molecular composition of C14H8Cl4. This metabolite corresponded to 1,1'-(2,2-dichloroethene-1,1-diyl)bis(4-chlorobenzene) (p,p'-DDE), which is the degradation product of p,p'-DDT and confirmed the involvement of the Lac enzyme in p,p'-DDT degradation. Further analysis showed the addition of (2H) resulting in the formation of a fragment ion at m/z 318 and a base peak of m/z 246, supporting the molecular composition of C14H8Cl4 and corresponding to 1,1-trichloro-2,2-bis(4-chlorophenyl)ethane (p,p'-DDD), indicating the transformation of DDT to DDE through dehydrohalogenation and subsequently to DDD via hydrogenation.

The degradation pathway also involved the subsequent dechlorination of p,p'-DDD, leading to the formation of 1,1'-(2-chloroethene-1,1-diyl)bis(4-chlorobenzene) (DDMU) with a molecular weight of 283. DDMU then underwent hydroxylation to form p,p'-DDOH (metabolite 5), facilitated by the hydroxyl radical oxidation of peroxidases (MnP or LiP). The oxidized compound, bis(4-chloro-3-hydroxyphenyl)acetaldehyde, generated by the action of AAO, exhibited an m/z of 267 and a molecular composition of C14H10Cl2O (metabolite 6). Further oxidation reactions potentially catalyzed by AAO converted metabolite 6 to bis(4-chlorophenyl)acetic acid (p,p'-DDA) (metabolite 7). The subsequent meta-ring cleavage by MnP and/or laccase through a hydroxyl radical mechanism led to the formation of single-ring aromatic compounds, including 1-(4-chlorophenyl)ethan-1-one (metabolite 8) with m/z 154, 4-chlorobenzaldehyde (metabolite 9) with m/z 140, and 4-methylbenzoic acid (metabolite 10) with m/z 136.

The degradation pathway of p,p'-DDT exhibits primarily multidirectional characteristics, as evidenced by the detection of two major hydroxyl derivatives. One of these derivatives, (metabolite 11) with a molecular composition of C14H8Cl4O and a mass-to-charge ratio (m/z) of 335, is identified as 2-chloro-5-[2,2-dichloro-1-(4-chlorophenyl)ethenyl]phenol. This hydroxyl derivative undergoes electron and H⁺ removal from the hydroxyl phenolic group through the action of peroxidases, resulting in the formation of a phenoxyl radical (Nadar et al. 2016b). Subsequent hydroxyl radical oxidation leads to the generation of a dihydroxylation product with an m/z of 352 and a molecular composition of C14H8Cl4O, identified as 3,3'-(2,2-dichloroethene-1,1-diyl) bis (6-chlorophenol), referred to as metabolite 12. Notably, peroxidase enzymes play a crucial role in initiating the oxidative dechlorination step during the degradation of various chlorinated phenols (Mohamed et al. 2023).

Enzymes produced extracellularly by white-rot fungi exhibit remarkable versatility in their ability to mineralize structurally diverse and highly persistent organic pollutants, which share similarities with lignin. Previous studies have demonstrated variations in the efficiency of enzymatic hydrolysis depending on the specific extracellular products interacting with different substrates (Wang et al. 2018). Peroxidases responsible for lignin degradation depend on a source of H₂O₂, which is supplied through the action of oxidases produced by fungi. Aryl-alcohol oxidases (AAOs, EC 1.1.3.7), also known as veratryl-alcohol oxidases, aromatic alcohol oxidases, or benzyl-alcohol oxidases, are a class of flavin-adenine-dinucleotide (FAD)-containing enzymes that catalyze the oxidation of aromatic and aliphatic allylic primary alcohols, concurrently reducing molecular oxygen to H₂O₂ (Urlacher and Koschorreck, 2021). The potential of AAOs to efficiently provide hydrogen peroxide for reactions catalyzed by peroxidases and peroxygenases is widely recognized. Additionally, several other flavin-containing oxidases, including vanillyl alcohol oxidase (EC 1.1.3.38) and 4-hydroxymandelate oxidase (decarboxylating; EC 1.1.3.19), possess the ability to oxidize aromatic or phenolic compounds (de Jong et al. 1992; Ewing et al. 2020; Martin et al. 2020). The mass spectra of identified by-products formed during the degradation process are presented in the supporting information (figures S1, S2, and S3).

Table 2

p,p'- DDT and identified p,p'- DDT transformation products detected by GC/MS during biodegradation by the immobilized enzymes biocatalyst
No.	Compound Name	RT (min)	Fragments (m/z)	MW	Formula
1	p,p'-DDT	36.98	199, 214, 235,246,354	354	C₁₄H₉Cl₅
2	p,p'-DDE	34.06	165, 176, 246, 318	316	C₁₄H₈Cl₄
3	p,p'-DDD	32.96	176, 246, 318, 320	318	C₁₄H₁₀Cl₄
4	p,p'DDMU	-------	-------	283	C₁₄H₉Cl₃
5	p,p'- DDOH	26.31	199, 213, 255,267	267	C₁₄H₁₀Cl₂O
6	bis(4-chloro-3-hydroxyphenyl)acetaldehyde	35.52	125, 246, 265	265	C₁₄H₁₀Cl₂O
7	p,p'-DDA	32.96	199,233, 264,281	281	C₁₄H₁₀Cl₂O₂
8	4-chlorobenzoic acid	7.04	91,121,136, 139, 154	154	C₈H₇ClO
9	4-chlorobenzaldehyde	7.02	91,121, 136, 140	140	C₇H₅ClO
10	4- methylbenzoic acid	15.76	91, 121, 136	136	C₈H₈O₂
11	2-chloro-5-[2,2-dichloro-1-(4-chlorophenyl)ethenyl]phenol	32.96	191, 228, 246, 334	334	C₁₄H₈Cl₄O
12	3,3'-(2,2- dichloroethene-1,1-diyl) bis (6-chlorophenol) .	28.74	123,149,222,264,352	350	C₁₄H₈Cl₄O₂

We have successfully created a stable and efficient enzyme biocatalyst by immobilizing multi-enzymes onto nano-silica. The immobilized enzymes demonstrated remarkable activity and stability across a wide pH range (4 to 9) and temperature range (20 to 55°C). This biocatalyst effectively degraded p,p'-DDT, completely eliminating it within a 12-hour incubation period at pH 5 and 30°C. The analysis also identified eleven major metabolites during the degradation process, indicating the effectiveness of the developed biocatalyst in treating environmental pollution caused by organic wastes. These findings contribute to the advancement of bioremediation technologies and have the potential to significantly impact environmental sustainability.

Acknowledgment

The authors acknowledge the Science and Technology Development Fund (STDF), Egypt (Project No. 30202) for supporting this work.

Conflict of interest statement

On behalf of all authors, the corresponding author states that there is no conflict of interest.

Author contribution

Tamer M. Salem: experiment and manuscript drafting

Nashwa A. H. Fetyan: experiment and manuscript drafting

Ahmed Aboueloyoun Taha: characterization, manuscript writing and proofreading

Farag M. Malhat: pesticide measurements

Amal Ibrahim Ramadan: experiment

Data Availability Statement

Data available upon request

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Covalent Immobilization of Multi-Enzyme Biocatalysts on Nano-Silica for Efficient p,p'-DDT Degradation

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Abstract

Figures

Introduction

Materials and Methods

2.1. Chemicals and media

2.2.1. Fungal strain and culture conditions

2.2.2. Purification of ligninolytic enzymes

2.2.3. Enzymatic activities and protein content

2.3.1. Synthesis of nano silica powder from rice husk:

2. 3.2. Covalent immobilization of oxidoreductase enzymes on nano-silica powder:

2.3.4. Effect of pH and temperature on the activity and stability of thr immobilized biocatalyst

2.3.5. Reusability assessment of the immobilized biocatalyst

2.4.1. Optimal reaction pH conditions:

2.4.2. Analysis and measurements

2.4.3 Extraction and determination of metabolites

Results and Discussion

3.1. Ligninolytic enzymes production:

3.2. Biochemical Characterization of nano-silica immobilized enzymes biocatalyst

3.3. Reusability assessment of the immobilized enzymes biocatalyst

3.4.1. p,p'- DDT degradation efficiency by the immobilized enzymes biocatalyst:

Conclusion

Declarations

References

Supplementary Files

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