In this work, the peroxidase enzyme was extracted from cabbage legs obtained from restaurant waste. Additionally, a comparison was made between crude and pure enzymes to evaluate their efficacy as catalysts for the biodegradation of common contaminants, textile dyes, and bisphenol A. The concentration of protein, bioactivity of peroxidase, and specific activity of both crude and pure enzymes were experimentally determined. The extracted peroxidase enzyme displayed optimal bioactivity at pH 6 and a temperature of 40°C. Moreover, it exhibited good stability across a wide pH range (3–7), retaining 65% of its original bioactivity at 60°C. The addition of FeSO4 and ZnCl2 enhanced enzyme activity by 153% and 120%, respectively, whereas exposure to CuCl2, AgNo3, and HgCl2 ions reduced enzyme activity by 60%, 53%, and 25%, respectively. The crude enzyme exhibited remarkable efficiency in decolorizing the synthetic dyes, with percentage decolorization of 86% and 78% for reactive blue 49 (RB) and reactive green 19 (RG), respectively, after a 10-hour incubation at the laboratory scale. Similarly, the pure enzyme exhibited a decolorization percentage of 79% and 75% for RB and RG, respectively, under the same conditions. The crude enzyme showed a high degradation efficiency of 92.8% for bisphenol A degradation in aqueous solution after 5 h.
Short Report
Extraction of peroxidase enzyme from cabbage legs for catalytic biodegradation of textile dyes and bisphenol A in wastewater: Purification and characterization technologies
https://doi.org/10.21203/rs.3.rs-3210800/v1
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Cabbage legs
Peroxidase catalyst enzyme
Purification
Characterization
Dyes and Bisphenol A degradation
Today's global emergency is the rise of water pollution. One of the biggest problems is the increased use of chemicals in many different industries. Large-scale application of chemical phenolic contaminants, such as reactive blue (RB), reactive green (RG), and bisphenol A (BPA), in several industries, such as textiles, leather, paper, and cosmetic, have been dramatically expanded because they are non-biodegradable in natural conditions. These hazardous contaminants can seriously damage the aquatic life and also exhibit dangerous impacts on the human health and ecosystem.
It has been established that some dyes and phenols provide a possible ecological danger due to their high toxicity, carcinogenicity, and/or mutagenicity (Bilal et al., 2016a, b; Skoronski et al., 2014). Bisphenol A (BPA) has received more attention in the environmental sector because it can disrupt physiological functions by affecting the endocrine system. However, small concentrations of BPA are undoubtedly released into the environment via manufacturing and processing technologies because BPA is a crucial material in industrial applications. The elimination of BPA is a vital issue owing to its impacts on human health resulting from cardiovascular disease, diabetes, cancer, and other health deficits (Venkatesan et al., 2015).
Currently, decontamination methods are either prohibitively expensive or harmful to health. Furthermore, numerous treatment techniques often result in the accumulation of secondary waste products. Therefore, new solutions for environmental cleanup employing green technology as a catalyst must be developed (Bilal and Asgher, 2015). Toxic pollution can be removed using a wide variety of physical and/or chemical-based procedures, but enzyme-based treatments provide a far safer option. Recent years have seen a rise in interest in catalyst enzyme -based biological treatments due to their many benefits, including their low environmental impact, high removal efficiency, high specificity, low maintenance requirements, high reproducibility, and broad contamination remediation potential. (Asgher et al., 2014).
Cabbage legs of plants are one of the major sources of peroxidase catalyst enzyme; they are cheap and widely available in local markets. The peroxidase enzyme (E.C. 1.11.1.7) has been demonstrated as an efficient, economic, and environmentally preferable catalyst for detoxification and decolorization of phenolic compounds in wastewater effluents (Sekuljica et al., 2015). Importantly, peroxidases can be categorized into three major classes based on metal-binding properties and amino acid homology. Class I includes cytochrome peroxidase and ascorbate peroxidase. Class II involves lignin-degrading peroxidases and encompasses manganese peroxidases. Class III secrete plant tissues, which have the capability to degrade various phenolic substances like catechin, chlorogenic acid, pyrogallol, catechol, and guaiacol (Pandey VP et al., 2017).
Moreover, peroxidase enzymes have been identified, isolated, sequenced, and described in various species, like bacteria, higher plants, and fungi (Zhang Z et al., 2018). The structural characteristics, kinetic properties, and substrate specificity of peroxidases make them effective in driving the bioreaction at extreme environmental conditions such as a wide range of pH and temperature, as well as high salinity and pollutant concentrations. Beyond that, peroxidases have the ability to catalyze the degradation process of resistant azo dyes in the existence of hydrogen peroxide, resulting in the formation of insoluble polymeric products that can be easily separated. Finally, enzymatic treatment offers a vital alternative to eliminate the delays associated with microbial acclimatization.
2.1. Chemicals and reagents
The cabbage waste (Brassica oleracea var.) was employed as a peroxidase catalyst enzyme, which was obtained from the local market for vegetables. The textile dyes, such as reactive blue 49 (RB) and reactive green 19 (RG) were made in Switzerland and provided by a textile general factory in Iraq, while bisphenol A was supplied from Hi-media/India. The dyes and bisphenol A detailed characteristics, along with their structures, are given in Table 1. Sodium acetate, potassium phosphate, Tris-HCl, pyrogallol, G-250 Coomassie Brilliant Blue, bovine serum albumin, and Sephadex G-150 were acquired from Sigma-Aldrich, USA. CuCl2, MgSo4, FeSO4, ZnCl2, AgNo3, and HgCl2 from CDH England, and acetonitrile as a mobile phase for HPLC (SYKAMN) Germany. The analytical-grade compounds utilized in this investigation were purchased from commercial sources. Before the experiments, deionized water was utilized to synthesis the wastewater stock of 40 mg/L of each dye and bisphenol A.
|
Dye Name |
λmax (nm) |
MW (g/mol) |
Structure |
|---|---|---|---|
|
Reactive Blue 49 |
595 |
882.19 |
|
|
Reactive Green 19 |
545 |
1418.94 |
|
|
Bisphenol A |
210 |
228.291 |
|
2.2. Methods
2.2.1. Extraction of plant peroxidase
The Cabbage legs were homogenized in a phosphate buffer solution with a concentration of 0.2 M (500 mL, pH 6.5) in order to obtain the crude enzyme extract (100 g). Afterward, the obtained solution was agitated at low speed (100 rpm) via a mechanical shaker (MRAD Corporation 311 series) of the Omni General Laboratory Homogenizer (GLH). Then, the solution was maintained at a temperature of 25°C for 24 hours, and the produced mixture was filtered through a double-layered cheesecloth to remove any solid particles and impurities, followed by centrifuging at 8000 rpm for 15 min using a Cole-Palmer VS-13000 instrument. After centrifugation, the obtained material was carefully gathered and preserved at a temperature of 10°C as the crude enzyme extract for further analysis and characterization (Tiwari, 2015).
2.2.2. Estimation of enzymatic activity
The hydrogen peroxide (H2O2) incorporated with the pyrogallol substrate was employed to assess the enzymatic activity of the peroxidase catalyst. The reaction involved the oxidation of pyrogallol, which was stimulated by the peroxidase catalyst, resulting in the formation of purpurogallin, an orange-brown product. Spectrophotometric analysis was employed to measure the concentration of purpurogallin (λ max = 420 nm) at a temperature of 25°C. The assay mixture companied of 0.2 mL of each of crude enzyme extract, 1% pyrogallol, 6% H2O2, and phosphate buffer (pH = 6.5). The rate of bioreaction was measured via a UV/Vis spectrophotometer (UV-9200, BIOTECH ENGINEERING) at a wavelength of 420 nm. Enzymatic activity was quantified in units (U), which represent the amount of enzyme able to catalyze and convert 1.0 mmol of H2O2/min at a temperature of 25°C and pH 6.5. Eq. (1) was applied to estimate the enzymatic activity of the peroxidase catalyst as follows (AL-Sa’ady et al., 2018):
$$Activity \left(\frac{\varvec{U}}{\varvec{m}\varvec{L}}\right)=\frac{\left(\frac{\varvec{d}\varvec{A}}{\varvec{d}\varvec{t}}\right)\times \varvec{A}\varvec{V}\times \varvec{D}\varvec{F}}{\varvec{E}\varvec{V}\times {\in }_{420} }$$
1
Where:
\(dA/dt\) : is the rate of increase in absorbance per minute in comparison to the flat part of the curve (min− 1)
\(AV:\) the total volume (mL)
\(DF\) : the factor of dilution
\(EV\) : the volume of enzyme extract (mL)
∈420: purpurogallin molar absorptivity at 420 nm (mM− 1cm− 1).
2.2.3 Total proteins estimation
The enzymatic activity assay involved estimating the protein content related to the raw enzyme extract. Bovine serum albumin (BSA) was used as a standard for protein quantification. To determine enzyme activity, protein concentration, and specific activity, the protein concentration was measured using the Bradford technique (Bradford, M. 1976). A standard curve for the Bradford assay was constructed using BSA. Triplicate preparations of successive concentrations ranging from 0.1 to 1.0 mg/ml were made from the stock solution of BSA. The resulting concentrations are presented in Table 2.
|
Tube Number |
Stock Solution Volume (BSA) (ml) |
Protein Concentration (BSA) (mg/ml) |
|---|---|---|
|
1 |
0.1 |
0.1 |
|
2 |
0.2 |
0.2 |
|
3 |
0.3 |
0.3 |
|
4 |
0.4 |
0.4 |
|
5 |
0.5 |
0.5 |
|
6 |
0.6 |
0.6 |
|
7 |
0.7 |
0.7 |
|
8 |
0.8 |
0.8 |
|
9 |
0.9 |
0.9 |
|
10 |
1 |
1 |
For each concentration of BSA, 0.1 mL was combined with Tris-HCL buffer (0.02 M and 0.4 mL), followed by adding Coomassie Brilliant Blue solution G250 (CBBG = 2.5 mL). The obtained mixture was thoroughly mixed and kept at room temperature. Besides, the blank solution was provided by blending CBBG (2.5 mL) with Tris-HCl buffer (0.5 mL). After a 5-minute incubation period, the absorbance of each concentration was detected at 595 nm. The correlation between protein absorbance and the corresponding protein concentration (mg/ml) was plotted and fitted by a standard curve. Please refer to Fig. 1 for the graphical representation of the standard curve.
2.2.3.1 Estimation the concentration of protein in the samples
In a test tube, 4 ml of Tris-HCl buffer was mixed with 0.1 ml of crude enzyme. After that, 2.5 ml of CBBG solution was dropped into each tube, and the attained samples were maintained for 5 min at room temperature. Afterward, 2.5 ml of CBBG solution was mixed with 0.5 ml of Tris-HCl buffer to prepare the blank solution. Then, the spectrophotometer reading for the absorbance of the colorimetric solutions was recorded at a wavelength of 595 nm. Finally, the protein concentrations in the unknown samples were measured, which represents the slope of the BSA plots.
2.2.3.2 Estimation of specific activity
Equation (2) was employed to estimate the specific activity of the peroxidase catalyst as follows (Whitaker and Bernard, 1972):
$$\varvec{S}\varvec{p}\varvec{e}\varvec{c}\varvec{i}\varvec{f}\varvec{i}\varvec{c} \varvec{a}\varvec{c}\varvec{t}\varvec{i}\varvec{v}\varvec{i}\varvec{t}\varvec{y} \left(\varvec{U}/\varvec{m}\varvec{g} \varvec{p}\varvec{r}\varvec{o}\varvec{t}\varvec{e}\varvec{i}\varvec{n}\right)= \frac{\varvec{E}\varvec{n}\varvec{z}\varvec{y}\varvec{m}\varvec{e} \varvec{a}\varvec{c}\varvec{t}\varvec{i}\varvec{v}\varvec{i}\varvec{t}\varvec{y} (\varvec{U}/\varvec{m}\varvec{l})}{\varvec{P}\varvec{r}\varvec{o}\varvec{t}\varvec{e}\varvec{i}\varvec{n} \varvec{c}\varvec{o}\varvec{n}\varvec{c}\varvec{e}\varvec{n}\varvec{t}\varvec{r}\varvec{a}\varvec{t}\varvec{i}\varvec{o}\varvec{n} (\varvec{m}\varvec{g}/\varvec{m}\varvec{l})}$$
2
2.2.4 Purification and Characterization of catalyst enzyme
The purification process was accomplished by passing the dialysate solution over a Sephadex G-150 column (50 cm × 2.5 cm) packed with phosphate buffer (0.2 M, pH = 6.5), followed by eluting via 500 mL of sodium phosphate buffer (0.2 M). After that, Peroxidase-active fractions were collected together, and the absorbance of the eluents was measured at 280 nm to determine the protein concentration.
2.2.4.1Optimum pH effect
The optimal pH range for both peroxidase activity and stability was determined by observing the enzyme's performance using the assay described in the previous section. A 3 ml solution of pyrogallol substrate was prepared using different buffers to achieve pH ranges of 4–5 (0.2M sodium acetate buffer), 6–7 (0.2M phosphate buffer), and 8–9 (0.2M Tris-HCl buffer). To each buffer, 0.3 ml of peroxidase enzyme was added, and the mixture was incubated for 30 minutes. Enzyme activities were then measured at 420 nm according to the method outlined by Al-Sa'ady et al. (2018). The optimal pH value for maximum bioactivity of peroxidase was specified by plotting the relationship between the bioactivity of the peroxidase catalyst and pH. Following the 30-minute incubation period under various pH conditions, the pH stability of the enzyme was examined by assessing the remaining activity at 420 nm. The percentage of remaining activity of the peroxidase enzyme was calculated, and a plot was generated to illustrate the relationship between pH and the percentage of remaining activity.
2.2.4.2 Optimum temperature effect
To estimate the optimum temperature for peroxidase activity, a 3 ml solution of pyrogallol substrate was mixed with 0.3 ml of peroxidase enzyme and incubated for 30 minutes. The incubation temperature ranged from 20 to 80°C. Peroxidase activities were then measured at 420 nm using a spectrophotometer, and the relationship between remaining activity and temperature was plotted based on the findings reported by Mizobutsi et al. (2010). For the temperature stability analysis, a water bath was utilized to incubate 0.3 ml of partially purified peroxidase for 30 minutes within the temperature range of 20 to 80°C. Immediately after incubation, the peroxidase enzyme test tubes were transferred to a cool water bath. The remaining enzyme activities were subsequently determined at 420 nm using the method described by AL-Sa'ady et al. (2018). A plot illustrating the relationship between temperatures and remaining activity (stability) was generated based on the results obtained.
2.2.4.3 Impacts of metal ions
The influence of inorganic ions on the bioactivity of enzymes was investigated through introducing 5 mM of CuCl2, MgSo4, FeSO4, ZnCl2, AgNo3, and HgCl2 with a volume ratio of 1:1 (v/v) (metal ions: enzyme) at a temperature 40˚C, incubation time of 15 min both with and without metal ions. The enzyme activity was estimated under standard assay conditions. Lastly, the residual bioactivities were estimated by taking the amount of controlled bioactivity to be 100%.
3.1. Application of cabbage leg peroxidase in synthetic dye decolorization
The WCP was employed to investigate the decolorization activity of synthetic dyes, in which the reactive blue 49 (RB) and reactive green 19 (RG) were selected as pollutant models with concentrations of 40 mg/L. The UV/Visible spectrophotometer was employed to analyze the prepared solutions to obtain the optimum wavelength of each dye. After adding buffer to each dye solution, the initial absorbance of RB and RG was measured. The effect of WCP on dye decolorization was studied under optimal reaction conditions, specifically using the defined WCP concentration and the optimum wavelengths. The degradation reaction of dyes was attained under the following conditions: 40 mg/L of dyes solution (5 mL), 0.5 mL of both crude and pure peroxidases, 0.5 mL of 2 Mm H2O2, and 2.5 mL of 0.2 M pH 6.5 phosphate buffer.
The bioreaction mixture was incubated at 40 ºC, and the values of final absorbance were recorded at 1, 5, and 10-hour intervals. The decolorization efficiency of synthesized catalyst was calculated using Eq. (3), allowing estimation of the extent of decolorization achieved during the experiment as follows in Eq. (3) (Takkar et al., 2021).
\(\varvec{P}\varvec{e}\varvec{r}\varvec{c}\varvec{e}\varvec{n}\varvec{t}\varvec{a}\varvec{g}\varvec{e} \varvec{D}\varvec{e}\varvec{c}\varvec{o}\varvec{l}\varvec{o}\varvec{r}\varvec{i}\varvec{z}\varvec{a}\varvec{t}\varvec{i}\varvec{o}\varvec{n}=\frac{\varvec{A}\varvec{i}-\varvec{A}\varvec{f}}{\varvec{A}\varvec{i}} \times\) 100% (3)
Where:
$$Ai = initial absorbance before decolorization$$
$$Af = final absorbance after incubation.$$
3.2. Application of cabbage leg peroxidase in Bisphenol A degradation
Bisphenol A (BPA) was employed as another pollutant model to reveal the biodegradation of the crude and pure catalyst WCP. High-Performance Chromatography-UV was used to determine the initial and final BPA concentrations. The standard concentration (40 mg/L) of BPA solution was prepared in the absence of any enzyme. The catalytic reactions were conducted in a 5 mL class tube as a batch reactor under the following operational conditions: crude and partially purified WCP volume of 0.5 mL (crude and partially purified), H2O2 concentration of 2 mM (particularly fixed), BPA concentration of 40 mg/L, and phosphate buffer concentration of 0.2 M (to maintain pH value of 6). Moreover, the bioreaction mixture was vigorously shaken and incubated at 40 ºC, and the final concentration values were collected after incubation at 1 h and 5 h intervals. As a result, the biodegradation efficiency could be estimated as follows:
\(\varvec{P}\varvec{e}\varvec{r}\varvec{c}\varvec{e}\varvec{n}\varvec{t}\varvec{a}\varvec{g}\varvec{e} \varvec{d}\varvec{e}\varvec{g}\varvec{r}\varvec{a}\varvec{d}\varvec{a}\varvec{t}\varvec{i}\varvec{o}\varvec{n}=\frac{\varvec{C}\varvec{i}-\varvec{C}\varvec{f}}{\varvec{C}\varvec{i}} \times\) 100% (4)
Where:
$$Ci = initial concentration before decolorization$$.
$$Cf = final concentration after incubation.$$
The purification technique for peroxidase from cabbage plant legs is given in Table 3. Due to their high polyphenol content, cabbage legs turn brown during the peroxidase extraction process. To get a uniform enzyme, the purification procedure used ultrafiltration concentration followed by sephadex G150 gel filtration chromatography purification. In Fig. 2, we see an example of an elution profile for a purification technique for cabbage legs. High peroxidase activity fractions (tubes 11–28) were employed in enzyme purification, characterization, and decolorization investigations because their peak correlated with the protein peak. The method employed increased specific activity by over 2.14-fold in comparison to crude extract, with a 39% yield, as shown in Table 3.
After the last chromatographic process, SDS-PAGE was employed to estimate the purity of the catalyst enzyme. In relation to the standard molecular weight, Fig. 3 displays gel filtration results showing a single band with a molecular weight of 55,000 Daltons. These peroxidases exhibit similar molecular weights to those found in horseradish and artichoke leaves, as well as in other food sources (Cardinali et al., 2007; Sakharov et al., 2000). The molecular weights of the peroxidase enzyme can range from 30,000 to 60,000 Daltons.
Table (3). Description of Cabbage Leg Peroxidase Purification.
|
Purification steps |
Volume (ml) |
Activity (U/ml) |
Protein concentration (mg/ml) |
Specific Activity (U/mg) |
Total Activity (U) |
Purification Fold |
Yield (%) |
|---|---|---|---|---|---|---|---|
|
Crude Enzyme |
100 |
4977 |
4.35 |
1144 |
497700 |
1 |
100 |
|
Crude with Sucrose Concentration |
28 |
7296 |
4.81 |
1516 |
204288 |
1.33 |
41 |
|
Gel filtration Sephadex G-150 |
46 |
4235 |
1.73 |
2448 |
194810 |
2.14 |
39 |
4.1 The characterization of purified peroxidase from waste cabbage.
4.1.1 Impact of pH.
The impact of pH on the enzyme bioactivity was examined, and the oucomes are summarized in Fig. 4a. The attained findings recorded desirable activity of partly purified WCP at pH values between 3 and 9, with the maximum activity received at pH 6.
The residual activity was incubated at room temperature in several buffers with different pH values ranging from pH (3–9) to investigate the pH value and stability of WCP. The partial peroxide was stable throughout a wide pH range (3–7) according to the results of the pH stability testing as shows in (Fig. 4b).
4.1.2. Effect of temperature
An optimal temperature of 40°C was revealed for the partly purified cabbage peroxidase, rising temperatures cause a dramatic rise in peroxidase activity, which at a temperature of 40°C reaches a peak. As the temperature rises over 40°C, peroxidase activity rapidly decreases, reaching minimum values between 70–80°C as shown in (Fig. 5-a).
To finding the thermal stability of purified WCP, the residual activity is achieved by incubation at PH 6 in a series of temperature values ranging from (20–80) ºC. According to the temperature stability experiment's findings, the partly purified peroxide was stable throughout a wide temperature range from (20–40) ºC as shown in (Fig. 5b).
4.1.3 Effect of metal ions
Table 4 displays the influence of metal ions on the bioactivity of peroxidase enzyme at a concentration of 5 Mm (1:1 (v/v) ratio), an incubation time of 15 min, and at 40°C. The peroxidase bioactivity was improved by FeSo4, ZnCl2, ions, with a remaining activity of 153% and 120%, respectively. Moreover, CuCl2, AgNo3, and HgCl2 ions retard the enzyme activity by 60%, 53%, and 25%, respectively, whereas with the metal ions of MgSo4, no change in cabbage peroxidase remaining activity was noticed with a percentage of 96%. Moreover, Pandey and Dwivedi (2011) found that divalent cations activate the peroxidase enzyme at metal concentrations of 50 mM but inhibit the enzyme at higher values.
|
Metal ion (5 mM) |
Relative activity (%) |
|---|---|
|
Control |
100 |
|
FeSo4 |
153 ± 3.3 |
|
ZnCl2 |
120 ± 3.1 |
|
CuCl2 |
60 ± 2.5 |
|
AgNo3 |
53 ± 2.1 |
|
HgCl2 MgSo4 |
25 ± 1.6 96 ± 3.5 |
4.1.4 Effect of reaction time on dyes decolorization
The amount of time that the dye is in touch with the absorbent during the color process of decomposition is one of the key aspects mediated by RB and RG dye degradation. The catalytic reaction contained 0.5 mL of WCP, 5 mL of RB and RG dyes (40 mg/L, pH 6), H2O2 concentration of 2 mM, and temperature of 40 ºC.
Figure 6 and Fig. 7 represents the degradation profiles as a function of time after 1,5, and10 h of incubation in a stainless-steel water bath MRC model HH-S2 at 40∘C. The results indicated that peroxidases rapidly degraded dyes in the first hours. This is because free enzymes have a several active sites accessible for random collision and interaction with substrate molecules.
The percentage decolourization after 10h for each dyes by crude enzyme centrifuge at 8000 rpm for 15 min was found to be 86%, 78% for RB and RG respectively and decolourization by pure enzyme was 79%,75% for the same dyes. (Subramaniam et al.,2010) reported after incubation at 50°C for 4 h, tannery effluent dye was found to be decolored 23% by Moringa charantia peroxidase. (Onder et al., 2011) showed by using horseradish after 5min of contact time at a temperature of 40∘C the decolourization was 90% of naphthol blue peroxidase. (Husain et al., 2010) recorded after 5 h incubation at 40∘C ,85% decolourization of dyes by using fenugreek peroxidase.
4.1.5 Effect of reaction time on Bisphenol degradation
The calibration curve in Fig. 8 was employed to measure the remaining BPA concentration after the biodegradation process over WCP. The peak of the curve represented the detention time used to calculate the area under the curve to determine the final concentration. 0.5 mL of WCP-catalyzed reactions containing 5 mL of (40 mg/L) Bisphenol A at pH 6 and 40 ºC in the presence of 2 Mm/L H2O2 were tested.
(Fig. 9a), (Fig. 9b), (Fig. 10a), and (Fig. 10b) reflect the absorption as a function of time after 1 h and 5 h of incubation in a stainless-steel water bath MRC model HH-S2 at 40°C. The results indicated that peroxidases rapidly degraded the Bisphenol in the first hours. This is because free enzymes have several active sites accessible for random collision. The percentage degradation after 1h and 5h incubation for Bisphenol by crude enzyme was found to be 59%, 92.8% respectively and the degradation by pure enzyme was 44%,71% respectively for the same period.
Waste cabbage legs were used to extract, partly purify, and analyze the biochemical characteristics of the peroxidase enzyme and tested to find the enzyme's ability to biodegrade synthetic dyes and Bisphenol A in aqueous solutions. The attained outcomes demonstrated that the bioactivity and stability of the peroxidase enzyme drop dramatically at both very acidic and very basic pH. The ionic state of the side chain of amino acids of the catalyst is typically affected by the pH, because the active site of the side chain of amino acids often affects heme-binding. This may explain why pH has an effect on the bioactivity and stability of the catalyst enzyme. Additionally, ionic alterations in the heme group may be the cause of a decline in stability and bioactivity at low and high pH values.
The waste cabbage Peroxidase (WCP) appears to be a thermostable catalyst enzyme, according to the results of thermal stability profiles, and the increase in the detected limit causes the enzyme to be denatured. Therefore, the thermal stability was suppressed after increase the reaction temperature.
Furthermore, the optimum bioreaction factors for WCP were employed to boost the degradation activity of dyes and BPA pollutants. An ecologically friendly and sustainable operation with an economic treatment process was attained by using cheap raw materials and applying cost-effective procedures for the extraction of the peroxidase enzyme. Besides, low amounts of chemicals and moderate reaction conditions were employed in this work. The result suggests that the crude peroxidase enzyme was very efficient in the degradation of both contaminants. Therefore, it is recommended to develop methods for producing organic enzymes in large-scale quantities.
Ethical Approval
Our study was approved by the Ethics Committee of environmental engineering department, college of engineering, university of Baghdad.
Consent to participate
Informed consent was obtained from all individual participants included in the study.
Consent to Publish
The authors affirm that human research participants provided informed consent for publication of the images
Author Contribution
Weam Abdulwahhab Mohammed (Methodology, Investigation, Software, and Writing-Original draft preparation), Mohanad J. M-Ridha (Data curation and supervision). All authors read and approved the final manuscript.
Funding
The authors declare that no funds, grants, or other support were received during the preparation of this manuscript.
Competing Interests
The authors declare no competing interests.
Data Availability
The data sets used and analyzed during the current study are available from the corresponding author upon reasonable request.
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