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Article

Antioxidants, Antimicrobial, and Anticancer Activities of Purified Chitinase of Talaromyces funiculosus Strain CBS 129594 Biosynthesized Using Crustacean Bio-Wastes

by
Hossam S. El-Beltagi
1,2,*,
Omima M. El-Mahdy
3,
Heba I. Mohamed
3,* and
Abeer E. El-Ansary
2
1
Agricultural Biotechnology Department, College of Agriculture and Food Sciences, King Faisal University, Al-Ahsa 31982, Saudi Arabia
2
Biochemistry Department, Faculty of Agriculture, Cairo University, Gamma St, Giza 12613, Egypt
3
Biological and Geological Sciences Department, Faculty of Education, Ain Shams University, Cairo 1575, Egypt
*
Authors to whom correspondence should be addressed.
Agronomy 2022, 12(11), 2818; https://doi.org/10.3390/agronomy12112818
Submission received: 17 October 2022 / Revised: 8 November 2022 / Accepted: 10 November 2022 / Published: 12 November 2022
(This article belongs to the Special Issue Biotechnology of Microorganisms in the Agriculture Environment)
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Abstract

:
Talaromyces funiculosus strain CBS 129594 was optimized to promote chitinase activity under solid state fermentation using crustacean bio-wastes. The aim of the study was to use purified chitinase as antioxidant, antimicrobial, and anticancer activities. The results showed that the maximum enzyme yield (2.98 ± 0.2 U/g substrate) was obtained at 1:2 crab shell chitin with the inoculation size (2.5 × 106 v/v) after seven days of incubation, pH 6.5, using 0.20% of soybean meal, malt extract, and yeast extract and 100% cane and beet molasses as supplementation. The enzyme was purified with an overall yield of 7.22 purification fold with a specific activity of 9.32 ± 0.3 U/mg protein. The molecular mass of the purified chitinase was 45 kDa. The highest chitinase activity was detected at pH 6.5 and 40 °C. The purified chitinase was activated by Ca2+, Cu2+, Na+, Mn2+, and Mg2+. On the other hand, the enzyme activity was inhibited in the presence of Hg2+, Ag2+, and Li+ at 10 mM, while Zn2+ and Co2+ caused no effect compared to media without any metals. The scavenging of 2.2-diphenyl-1-picrylhydrazyl (DPPH) radicals and 2.2-pheny-l-1-bis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) increased with increasing the concentrations of the purified chitinase enzyme (100, 200, 300, and 400 µg/mL) which ranged from 48.7% to 57.8% and 8.87% to 63.73%, respectively. The IC50 value of DPPH radicals and ABTS of purified chitinase produced by T. funiculosus strain CBS 129594 was 199 and 306 μg/mL concentration, respectively. The purified chitinase inhibited the growth of Gram-negative bacteria (Pseudomonas aeruginosa, Escherichia coli), Gram-positive bacteria (Bacillus subtilis, Staphylococcus aureus), and fungi (Aspergillus niger, Candida albicans). The highest concentrations of purified chitinase (1000 µg/mL) caused the higher toxicity of cancer cell line MCF7 (97%), HCT116 (88.2%), and HepG2 (97.1%). In conclusion, we can conclude that chitinase can be produced from marine waste and can be used as an antioxidant, antibacterial activity, cancer therapy, and ecofriendly biocontrol agent.

1. Introduction

Seafood is described as freshwater and marine fish that are consumed in human diets [1]. Arbia et al. [2] noted that byproducts of seafood consumption, particularly shrimp shells, account for approximately 40–50% of total mass, and these wastes pose a significant environmental hazard because they break down progressively and take a lot of time. Every year, the seafood processing industries create around 6–8 million tons of crab, shrimp, and lobster shells globally [3]. Crustacean bio-wastes, particularly shrimp and crab shells, contain the greatest amount of chitin of any natural resource [4]. The shell waste is primarily composed of protein (30–40%), minerals (30–50%), and chitin (20–30%) [5].
Chitin (β-(1–4)-N-acetyl-d-glucosamine) is a cellulose-like biopolymer produced by many species of microorganisms. Chitin can be present in the exoskeletons of invertebrates, crustaceans, actinomycetes, arthropods, and insects, as well as in fungi, bacteria, and yeast cell walls [6]. At least 1013 kg of chitin is created and degraded each year in the biosphere. After cellulose, chitin is the second most prevalent natural polymer in the world. Chitin is insoluble in many solvents because of its rigid structure [7]. Chitin contains a high amount of nitrogen (6.89%), making it an effective chelating agent [8]. Chitin is catabolized in two phases, beginning with chitinases cleaving the chitin polymer into chitin oligosaccharides and continuing with chitobiases cleaving into N-acetylglucosamine and monosaccharides [9].
Chitinases (EC 3.2.1.14) are hydrolytic enzymes that cleave glycosidic linkages in chitin. As a result, chitinases have molecular weights ranging from 20 to 90 kDa [10]. Chitinases can break down chitin directly into low molecular weight chitooligomers, which have a variety of commercial, agricultural, and medical applications, including elicitor activity and anticancer potential [11,12]. Chitinases are able to bioconvert chitin into valuable chemicals that can be utilized in biotechnology, waste treatment, pharmacology, biomedicine, single-cell protein, separation of protoplasts from fungi and yeast, drug carriers, and the enzyme industry [13]. Furthermore, chitinase plays an important role as a biocontrol agent against various fungal diseases [14,15], as well as being used as a dietary supplement [16]. The manufacturing of extracellular chitinase is gaining popularity around the world and can be improved by changing the mix of the culture medium and the fermentation conditions [17]. Improved manufacturing conditions are crucial for achieving optimal output, productivity, and cost savings [18]. Furthermore, fungal chitinases play a significant role in the biocontrol of pests that attack various plants, causing economic losses around the world, reducing or replacing the use of chemical insecticides, and reducing the adverse environmental impacts [19].
The chitinase lytic enzymes released by the antagonists, breakdown the fungal cell wall, which is made up of chitin and glucan in addition to wall proteins, individually or collectively contributing to the biocontrol activity [20]. Additionally, this enzyme has a negative impact on the pathogen’s conidial germination, germ tube elongation, and may harm the oospores [20]. Cellular deformities, protoplasmic damages, mycelial distortion and lyses, leakage of cellular contents, and alterations in membrane permeability are a few more negative impacts of chitinase activity on pathogens [21]. Therefore, one of the primary mechanisms responsible for the biocontrol effects of chitinase and other lytic enzymes released by microbial antagonists is the loss of fungal cytoplasm caused by the enzymatic cell wall disintegration [21]. Wherever chitin is present in the host, chitinases are critical players in bacterial pathogenesis [22].
Cancer is a leading cause of death in the world’s population. According to the American Cancer Society, cancer will overtake cardiovascular disease as the second greatest cause of death in a few years [23]. Furthermore, chemotherapy has caused a major public health issue and, in some cases, had no effect on cancer cells [24]. This problem is exacerbated in economically developing countries due to the absence of accessibility to standard diagnostic services and the high price of therapy [25]. Free radicals are known to be a major cause of a variety of diseases, including heart disease and cancer [26,27,28,29,30,31]. Although oxidation is responsible for the production of energy in living organisms, excessive free radical formation and a lack of antioxidant defense play a part in oxidative damage, which is harmful to cells and is also linked to cancer formation due to its effect on DNA [32]. Because of the harmful impact of free oxygen radicals, numerous biological methods are involved in their elimination, including the scavenging of reactive oxygen species, chelating by transition metal catalysts, and using antioxidants to detoxify ROS [33,34,35,36]. Synthetic antioxidative agents with high radical scavenging activity have been linked to serious adverse effects [37]. Natural antioxidants are therefore recommended, and recent studies have shown that they can be produced by microbes in addition to plants, which are a great source of antioxidants [38,39,40].
In this regard, the current study aimed to use crustacean shell waste for the production of chitinase using Talaromyces funiculosus strain CBS 129594 via solid-state fermentation and study its applications and roles as antimicrobial, antioxidant, and anticancer potential.

2. Materials and Methods

2.1. Microorganisms

The screening study was done using a modified Czapek’s medium which contained chitin obtained from Sigma Aldrich (St. Louis, MO, USA) as a carbon and nitrogen source (0.5 g/50 mL media). A triple set of 250 mL conical flasks with 50 mL of the basal solution was prepared for each treatment, sterilized at 121 °C for 20 min under 1.5 atmospheric pressure, and then stored at room temperature. After the sterilization of the media, they were inoculated with a prepared spore suspension (105 spores/mL) of the tested ten fungi (Alternaria citri, Aspergillus clavatus, Aspergillus flavus, Fusarium oxysporum, Fusarium solani, Mucor circinelloides, Penicillum chrysogenum, Rhizopus stolonifera, Trichoderma viride, and Talaromyces funiculosus) and incubated at 30 °C for 7 days. The chitinase activity was determined in the filtrates of the ten fungi used. The amount of reducing sugars (N-acetylglucosamine GlcNAc) in the supernatant was used to determine chitinase activity according to Miller [41]. After screening of microorganisms, the fungus which produced high amounts of chitinase enzyme was used in the study.

2.2. Preparation of the Crustacean Shell Powder and Colloidal Chitin

Shrimp, crayfish, and crab shells were gathered and stored in a freezer before being transferred to the laboratory. The shells were rinsed with water, dried, and crushed to powder (800–1000 mm). Colloidal chitin was obtained from Sigma-Aldrich chitin (C7170 practical grade, powder) as described by Halder et al. [42] protocol.

2.3. Genetic Identification of the Fungus

The selected fungus used in this study was identified according to genetic identification. By biosynthesizing the major compound, 16S rRNA analysis was performed on the fungus. Fungal DNA was extracted in the lab using the Qiagen DNeasy Mini Kit protocol. The extracted DNA was subjected to a polymerase chain reaction using universal primers (TGCGGAAGGATCATTACCGAGTGCGGGCCCTCGGCCCAACCTCCCCCTTGTCTCTA) in Seoul, South Korea, at Macrogen Companies. Amplified DNA was sequenced, and the resulting DNA sequence was identical to the DNA sequence available at NCBI GenBank*. The gene sequence that resulted was submitted to the NCBI GenBank database and given an accession number (MH865441.12).

2.4. Chitinase Production through Solid State Fermentation (SSF) and Extraction

Czapek’s basal medium was used in the production of chitinase enzyme, but carbon and nitrogen sources were replaced separately by adding 5.0 g of the different natural substrates namely, shrimp chitin, crayfish chitin, crab chitin, and chitin of Sigma, and the pH was adjusted to 7.0 and the moisture became 70% [43]. After autoclaving, each flask was injected with T. funiculosus strain CBS 129594 (0.97–109 CFU/gds), mixed thoroughly, and incubated at 30 °C for 7 days. After shaking the solid ferments for 1 h at 120 rpm in a rotary shaker at 30 °C, the enzyme was dissolved in sterile distilled water (solid: water; 1:10). The solution was used as a crude enzyme source.

2.5. Determination of Chitinase and Protein

Chitinase activity was measured using 3,5-dinitrosalicylic acid (DNS) reagent, and chitin as substrate as described by Halder et al. [43]. The protein content was determined in each step of purification using Folin–Ciocâlteu phenol reagent as described by Lowry et al. [44] method and using bovine serum albumin as a standard.

2.6. Effect of Additives and Agro-Industrial on Chitinase Production

Different additives such as soybean meal, malt extract, and yeast extract were added at different concentrations to the media (0.05–0.10, 0.15–0.20, 0.25% w/v) to study the effect of these additives on the production of chitinase. Agro-industrial product such as cane and beet molasses were prepared as following:
A measure of 200 mL of either cane or beet molasses was diluted to 1.0 L with tap water, sterilized in the autoclave at 121 °C for 20 min at 1.5 atmospheric pressure kept at room temperature, and then filtered to remove the precipitated mud. The filtrate was kept in the refrigerator in dark bottles. Cane and beet molasses (sugar industry) were collected from sugar factories in Hawamdiah and Kafr-El-Sheikh, respectively.

2.7. Effect of Different Incubation Period, Inoculum Size, and NaCl on Chitinase Production

The effect of the incubation period was investigated at intervals ranging from 3–10 days and then the chitinase activity was measured. The effect of inoculum size was studied at the range of 1.9 × 106, 2.5 × 106, 4.7 × 106, 5.8 × 106, and 7.2 × 106 and the chitinase activity was measured to determine the optimum size. Moreover, in this study, the media was incubated with different NaCl levels (0.5–3.0%) for 24 h at 4 °C, and the chitinase activity was determined.

2.8. Purification of Chitinase Enzyme

To precipitate the proteins, the crude enzyme was mixed with 60% saturated ammonium sulphate and then centrifuged at 10,000× g for 15 min. The precipitate was dialyzed for 24 h after being solubilized in phosphate buffer (pH 7.0, 50 mM). The dialysate was loaded onto a column of diethyl-aminoethyl-cellulose (2 cm × 60 cm) (DEAE-cellulose was purchased from Sigma (St. Louis, MO, USA) and activated as per manufacturer’s instructions ). The proteins were dissolved with a linear gradient of 0–1.0 M NaCl containing the same buffer at a flow rate of 1 mL/min. An auto-fraction collector was used to collect fractions of 2 mL. Before being eluted with phosphate buffer, the chitinase-active fractions were concentrated and loaded onto a Sephadex-G100 (Sigma, St. Louis, MO, USA) gel filtration column (2–70 cm) (pH 7.0, 50 mM). At a flow rate of 0.5 mL/min, fractions of 2 mL were gathered and the chitinase activity was measured. Chitinase-active fractions were lyophilized and stored at −20 °C.

2.9. Molecular Weight Determination

A standard marker protein and 12% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was used to determine the molecular mass of purified chitinase. The gel was put in 0.25% Coomassie Brilliant Blue R-250 to stain it and then put overnight with acetic acid-methyl alcohol-water (5:5:1 v/v) to remove the excess stains.

2.10. Amino Acid Analysis

Samples of 1 mL of pure enzyme sample and 1 mg of protein were hydrolyzed by heating it for 24 h at 110 °C in a sealed, evacuated tube with HCl (6 N, 5 mL), and β-mercaptoethanol (10 mL). The excess HCl was removed by vacuum evaporation to dryness. Beckman Amino Acid Analyzer was used to determine amino acids in the sample [45].

2.11. Temperature, pH Optima, and Stability

Different temperatures (30–100 °C) and pH (4.0–8.0) were tested to select the optimum temperature, pH, and the thermal and pH stability of the purified chitinase was measured.

2.12. Effect of Metal Ions on the Purified Chitinase

The metal ions were pre-incubated for 1 h at optimum assay conditions with solutions of the followings:
Na+, Mg2+, Mn2+, Hg2+, Zn2+, Ca2+, Co2+, Cu2+, Ag2+, Li+ at final concentrations of 5 and 10 mM.
Surfactants: SDS and Tween-80 (5 and 10 mM).
Chelating agents: ethylene diamine tetra acetic acid (EDTA) at final concentrations of 5 and 10 mM.
Inhibitors: β-mercaptoethanol, phenylmethylsulfonyl fluoride (PMSF) at final concentrations of 5 and 10 mM.
In all cases, the residual chitinase activity was measured.

2.13. Biological Activities of the Crude Enzyme

2.13.1. Free Radical Scavenging Activity

The antioxidant activity of the crude chitinase enzyme was determined according to the method of Park et al. [46]. The purified enzyme stock ethanolic solution was diluted with ethanol to final concentrations ranging from 100–400 µg/mL enzyme. Then, one mL of DPPH (0.2 mM) was stirred continuously and incubated at room temperature in the dark for 30 min. The optical density was read at 517 nm using a spectrophotometer.
% DPPH = (Absorbance of Control − Absorbance of Sample)/(Absorbance of Control) × 100.
To calculate the IC50 value, the percentage of radical scavenging activity was charted against the corresponding concentration of the crude enzyme. The IC50 value is the maximum concentration of a compound required to inhibit it by 50%.

2.13.2. ABTS Radical Cation Decolorization Assay

A modified method by Re et al. [47] was used for the ABTS radical cation decolorization assay. First, 5 mL of 7 mM ABTS was added to an equal volume of 4.9 mM potassium persulfate. The mixture was kept at room temperature in the dark for 16 h and then 1.8 mL of the ABTS reagents were mixed with 0.2 mL of different concentrations of ethanolic extract of purified enzyme (100–400 µg/mL), and then the optical density was read at 734 nm. L-ascorbic acid (0.3 mM) was used as a control.
% Inhibition = (ABS control − ABS Sample)/(ABS control) × 100

2.13.3. Antimicrobial Activity of Pure Chitinase

The antimicrobial activity of the tested crude T. funiculosus strain CBS 129594 chitinase was carried out against Gram-positive (B. Subtilis, S. aureus), Gram-negative bacteria (P. aeruginosa, E. coli) and fungi (A. niger, C. albicans). Antimicrobial activity was measured by the well diffusion method as described by Espinel-Ingroff et al. [48,49] with slight modification. Measurement of the inhibition zones was done by placing 100 µL of the crude preparation separately, each one in 6 mm diameter wells cut in nutrient agar plates seeded by test bacteria and potato dextrose agar plates seeded by test fungi. After that, the plates containing bacteria were incubated at 37 °C for 24 h and the plates containing fungi were incubated at 30 °C for 72 h, and the diameter of zone inhibition was measured in mm.

2.14. Anticancer Activity

Determination of Sample Cytotoxicity on Cells (MTT Protocol)

Three different cell lines (colorectal carcinoma colon cancer (HCT116), breast cancer of the mammary gland (MCF7), and hepatocellular cancer (HepG2)) were used in this study. The cell lines were obtained and activated according to the ethics of the scientific community by VACSERA-Cell Culture Unit, Cairo, Egypt. (1) To create a full monolayer sheet, 1 × 105 cells/mL (100 L/well) of cells were added to the 96-well tissue culture plate. The plate was then incubated at 37 °C for 24 h. (2) After the cell monolayer had been washed twice with wash media, the growth medium was decanted from the 96-well microtiter plates. (3) The tested material was diluted twice in RPMI medium with 2% serum (maintenance medium). (4) Three wells served as the control wells and received only maintenance medium while 0.1 mL of each dilution was examined in different wells. (5) The 37 °C-incubated plate was inspected. The physical characteristics of toxicity, such as partial or total loss of the monolayer, rounding, shrinkage, or cell granulation, were examined in the cells. (6) A MTT solution (5 mg/mL in PBS) (BIO BASIC CANADA INC, Markham, Canada) was made and each well received 20 µL of the MTT solution. For five minutes, shake at 150 rpm to properly blend the MTT into the media and allow the MTT to metabolize for 1 to 5 h in an incubator (37 °C, 5% CO2). (7) Dump off the media (dry the plate on paper towels to remove residue if necessary) and resolve 200 µL of formazan (a metabolic product of MTT) in DMSO. To thoroughly combine the formazan and solvent, place on a shaking table and shake at 150 rpm for 5 min. Read the optical density at 560 nm and subtract the background at 620 nm as described by Mosmann [50] and Wilson [51]. The relative cell viability (RCV) in percentage was determined according to the following equation:
% RCV = (A570 of treated samples)/(A570 of untreated sample) × 100

2.15. Statistical Analysis

Each value was expressed as the mean of three independent experiments. Data were assessed by analysis of variance (ANOVA) through Duncan’s multiple range tests using SPSS software (SAS Institute Inc., Cary, NC, USA).

3. Results

3.1. Screening for Chitinase Production by Some Fungi

The screening of ten fungi (Alternaria citri, Aspergillus clavatus, Aspergillus flavus, Fusarium oxysporum, Fusarium solani, Mucor circinelloides, Penicillum chrysogenum, Rhizopus stolonifera, Trichoderma viride, and T. funiculosus) for the production of chitinase enzymes in shaken cultures lasting for 3–7 days. T. funiculosus produced a higher activity of chitinase about 1.4 ± 0.1 U/mL. As a result, this fungus was chosen for further investigation using solid-state fermentation (SSF), and it was molecularly identified using 16S rRNA.

3.2. Molecular Identification of the Fungus Producing the Most Chitinase

The 16S rRNA gene sequence was utilized to detect and contrast with other discovered sequences in the Gene Bank database using BLAST to show score similarities and calculate the statistical difference of matches. The results found a very close similarity with T. funiculosus strain CBS 129594 using the 16S rRNA gene sequence with 100% homology of the isolate CBS 129594. The phylogenetic analysis and tree were composed using the neighboring method (Figure 1 A–C). Based on the analysis of the DNA sequence, the isolated strain was identified as T. funiculosus strain CBS 129594 and deposited in GenBank with an accession no. MH865441.1.

3.3. Effect of Moisture Content (MC) and Shellfish Waste as a Substrate Medium Ratio on the Production of Chitinase on SSF Technique

The moisture content of the medium in solid substrate fermentation is very important for the growth of microorganisms, the production of enzymes, and enzyme activity. In the screening experiment, dry-milled substrates of some crustacean wastes such as shrimp chitin, crayfish chitin, crab chitin, and chitin purchased from Sigma were utilized to select the most effective for chitinase activity as the sole C/N source. The results in Table 1 revealed that chitinous wastes increased the chitinase activity because they are considered a good source of carbon and nitrogen. In comparison to the other shellfish wastes, the chitin obtained from sigma and crab shell chitin had the highest chitinase activity at all substrate and volume ratios.

3.4. Effect of Weight of Crab Shellfish Waste on Chitinase Production

The weight of crab shellfish waste has an effect on the production of chitinase. Different weights of crab shellfish waste (5, 10, 15, 20, and 25 g/250 mL conical flask) were used to evaluate the production of chitinase. The results showed that 10 g/250 mL conical flask gave the highest production of chitinase (3.24 ± 0.2 U/g). The production of chitinase decreased significantly with an increase in the weight of crab shellfish waste (Figure 2).

3.5. Optimization of Chitinase Production

Microbial enzyme biosynthesis is regulated by cultural and environmental factors. In this regard, a series of studies were conducted to determine the influence of those factors on the enzyme productivity by T. funiculosus strain CBS 129594 in order to identify the suitable factors to obtain the highest chitinase production under SSF. The production of chitinase increased with an increasing incubation period for some days (Figure 3A). The results showed that after 7 days of incubation, the highest chitinase yield (3.56 ± 0.04 U/g) was obtained, and after this period, the production of chitinase began to decrease. In addition, the production of chitinase by T. funiculosus strain CBS 129594 was affected by inoculum size (Figure 3B). The inoculation size (2.5 × 106 v/v) produces the highest amounts of chitinase, approximately 3.94 ± 0.2 U/g, while increasing the inoculation size caused a reduction in chitinase production.
Figure 4 illustrates the impact of the addition of various agro-industrial wastes and some additives to the production media on the production of the chitinase enzyme. The additives of soybean meal, malt extract, and yeast extract substantially increased chitinase production. The chitinase production significantly increased with increasing the percentage of soybean meal, malt extract, and yeast extract (0.05, 0.10, 0.15, 0.20, and 0.25%) except at 0.25%, there is a non-significant effect between soybean meal and malt extract. Furthermore, when 0.20% soybean meal, malt extract, and yeast extract were added to the production media, chitinase production increased to approximately 7.74 ± 0.2, 8.09 ± 0.4, and 8.99 ± 0.3 U/g, respectively. In comparison to the other additives tested, the media containing yeast extract yielded the highest chitinase production.
The results on the influence of adding different concentrations of agro-industrials such as cane and beet molasses (sugar industry) (1:3, 1:1, 3:1 cane molasses, beet molasses, and 100% molasses on chitinase production are presented in Figure 5. All concentrations of sugar cane increased the chitinase production as compared to control media without any supplementation. The addition of 100% cane and beet molasses gave the highest chitinase production by about 9.78 ± 0.4 and 11.56 ± 0.5 U/g, respectively.
The results in Figure 6 show that the different concentrations of NaCl (0.5–3.0%) had a significant effect on T. funiculosus strain CBS 129594 chitinase production. The highest production of chitinase at 2% NaCl did not show any significant effect as compared to the control, while the other concentrations of NaCl caused a considerable reduction in chitinase production.

3.6. Chitinase Purification

The chitinase crude extract produced from T. funiculosus strain CBS 129594 was precipitated using 50–90% by (NH4)2SO4 precipitation. Table 2 shows the amount of protein, enzyme activity (U/mg protein), and specific activity values calculated for the fractions produced during the purification steps of the chitinase enzyme obtained from T. funiculosus strain CBS 129594. The chitinase enzyme was synthesized from T. funiculosus strain CBS 129594 and purified in two steps. The enzyme produced from T. funiculosus strain CBS 129594 using % (NH4)2SO4 saturation which has 1.93-fold purification, and 73.3% recovery was then injected into a Sephadex G-100 column. The total protein and chitinase activity recovered from Sephadex G-100 achieved approximately 65.4% and 2.89-fold purification of the application sample. The most effective fractions (13–19) from the Sephadex G-100 gel filtration column were transferred to an ion-exchange chromatography column containing DEAE-cellulose. The fractions (17–22) had the highest specific activity of 9.32 U/mg protein, 7.22-fold purification, and 60.8% recovery.

3.7. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE)

The results demonstrated that SDS-PAGE electrophoresis of pure chitinase revealed the existence of a specific protein band that has a molecular weight of 45 kDa (Figure 7).

3.8. Amino Acid of Purified Chitinase

The data in Figure 8 show that there are sixteen amino acid compounds observed in the purified chitinase enzyme. Aspartic acid was detected as the highest amino acid value (15.2%), followed by glutamic acid (12.3%), while histidine (2.1%) and methionine (1.6%) were found to be the lowest amino acid value.

3.9. Effect of Temperature and Thermal Stability on the Activity of Purified Chitinase

One of the important conditions for the efficacy of enzyme activities is reaction temperature. The optimum temperature for chitinase activity was observed to be 40 °C (12.36 ± 0.5 U/mL). As a result, the activity of chitinase decreased progressively as the temperature rose or fell above 40 °C (Figure 9A). At 70 °C, the relative stability of chitinase activity was 85.9% (Figure 9B).

3.10. Effect of pH on the Activity of Purified Chitinase

The yield of chitinase synthesized from T. funiculosus strain CBS 129594 was studied in relation to the initial pH of the fermentation medium. The highest chitinase activity was identified at pH 6.5, with 13.93 ± 0.9 U/mL, while the lowest chitinase activity was found at pH 8.0, with 8.01 ± 0.5 U/mL (Figure 10A). Furthermore, 90.6% of the relative stability of chitinase activity was identified at pH 6.5 (Figure 10B).

3.11. Effect of Metal Ions and Inhibitors on Purified Chitinase Activity

In a variety of enzyme-catalyzed processes, metal ions can act as inducers or inhibitors. The effects of various metal ions on chitinase synthesis were determined (Table 3). Ca2+, Cu2+, Na+, Mn2+, and Mg2+ were shown to be powerful activators, increasing enzyme activity by approximately 150%, 130%, 120%, 115%, and 110%, respectively, at 5 mM. In contrast, Hg2+, Ag2+, and Li+ at 10 mM decreased enzyme activity, although Zn2+ and Co2+ had no effect when compared to the control media without any metals. The effect of several enzyme inhibitors on T. funiculosus strain CBS 129594 chitinase activity was also investigated. Purified chitinase was stable and mildly more active at a 5 mM concentration of sodium dodecyl sulphate (SDS) (95%), Tween-80 (95%) as surfactants, EDTA (92%) as chelating agents, and PMSF (93%) as inhibitors.

3.12. Antioxidant Activity of Purified Chitinase

The ability of antioxidants to donate hydrogen was related to their influence on DPPH radical scavenging and ABTS. DPPH is a stable free radical that receives an electron or hydrogen radical to produce a stable molecule. The antioxidant capacity of pure chitinase synthesized by T. funiculosus strain CBS 129594 was investigated (Table 4). Table 4 shows that increasing the concentrations of the crude chitinase enzyme (100, 200, 300, and 400 µg/mL) increased the scavenging of DPPH radicals by about 48.7% to 57.8% and ABTS by about 8.87% to 63.73% as compared to ascorbic acid and butylated hydroxytoluene as control. The IC50 value of DPPH radicals and ABTS of purified chitinase produced by T. funiculosus strain CBS 129594 were 199 and 306 μg/mL concentration, respectively. The IC50 value means the concentration of the test sample required to suppress 50% of the free radicals. The high concentration of purified enzymes (400 μg/mL) caused the maximum inhibition percentage of DPPH and ABTS to be about 57.8% and 63.7%, respectively (Table 4).

3.13. Antimicrobial Activity of Purified Chitinase

Data in Table 5 illustrate the antimicrobial activity of the purified chitinase from T. funiculosus strain CBS 129594 against a set of microorganisms comprising Gram-negative bacteria (P. aeruginosa, E. coli), Gram-positive bacteria (S. aureus, B. subtilis), and fungi (C. albicans, A. niger). Purified chitinase has antimicrobial activity against all microorganisms used by using agar diffusion. The results revealed that the purified chitinase exhibited antimicrobial activity against P. aeruginosa, B. subtilis, A. niger, C. albicans, and E. coli with the inhibition zone diameters of about 38, 29, 26, 25, and 24 mm, respectively, compared to S. aureus (20 mm) (Figure 11 A–F).

3.14. Anticancer Activity of Purified Chitinase

This study revealed that different concentrations of purified chitinase (1000, 500, 250, 125, 62.5, and 31.25 µg/mL) affect the cell viability and toxicity of three cancer cell lines. The highest concentrations of purified chitinase (1000 µg/mL) caused the higher toxicity of cancer cell line MCF7 (97%), HCT116 (88.2%), and HepG2 (97.1%). The MTT assay revealed that the purified chitinase from T. funiculosus strain CBS 129594 had a highly toxic effect against the breast cancer of the mammary gland (MCF7) cells with an IC50 value of 101.81μg/mL and against hepatocellular cancer (HepG2) with an IC50 value of 118.74 μg/mL and moderate cytotoxic activity against colorectal carcinoma colon cancer (HCT116) with an IC50 value of 411.45 μg/mL (Table 6 and Figure 12 and Figure 13A–C).

4. Discussion

There has been a great deal of interest in the synthesis of microbial chitinases in recent decades, and microorganisms that synthesize a complex of mycolytic enzymes are thought to be potential biological control agents [52]. Furthermore, fungal species produce a wide range of chitinase isomers with varying catalytic characteristics [53]. In this study, ten fungi were used for screening in the production of chitinase. T. funiculosus produced a higher activity of chitinase about 1.4 ± 0.1 U/mL. These findings are in accordance with those of Duo-Chuan et al. [54], who observed that Talaromyces flavus CGMCC 3.4301 was able to synthesize chitinases when cultivated in the existence of chitin. The results of 16S rRNA confirmed a very similar relationship with T. funiculosus strain CBS 129594 using the 16S rRNA gene sequence with 100% homology of the isolate. The conversion of chitinous wastes such as crab, shrimp, and crayfish shells to be important products was considered an effective way of waste management and decreased environmental issues. In this study, crab shells produced the most chitinase when compared to other wastes using the solid-state fermentation (SSF) technique. In this regard, SSF is superior to submerged fermentation (SmF) in terms of high production, simple way, cost, energy savings, less effluent problem, and product stability because of less dilution, and is thus regarded as a viable method for industrial manufacture of a wide variety of value-added products [55].
The moisture content of the medium in SSF is critical for microorganism growth, enzyme synthesis, and enzyme activity [56]. Chitin sigma and crab shells chitin gave the highest moisture content and chitinase activity at all ratios of substrate and volume of media as compared to the other shellfish wastes. The results showed that 10 g/250 mL conical flask gave the highest production of chitinase (3.24 ± 0.2 U/g). Farag and Al-Nusarie [57] investigated the effect of chitin concentration on chitinase biosynthesis, and they found that when A. terreus was cultivated with shrimp-shell powder chitin at concentrations ranging from 5 to 20 g/L, chitinase content increased, while at concentrations greater than 25 g/L, the chitinase production reduced. Chitinase synthesis reduced as the weight of crab shellfish waste increased. The addition of a high shrimp-shell powder chitin in the culture medium for extended durations of incubation greatly inhibited chitinase synthesis [57]. A high shrimp-shell powder chitin concentration in the medium causes higher viscosity, which reduces oxygen availability and inhibits fungal growth and enzymatic activity. Similar results were reported by Shalaby et al. [58] who found that in all cases, chitin powder was the most favorable by Streptomyces halstedii H2 and led to the highest chitinase enzyme production (49.65 U/reaction), followed by crab shells powder, crab shells pieces, prawn shells, shrimp shells, and mollusk valves powder (44.52, 31.28, 26.24, 25.40, and 20.68 U/reaction, respectively).
The incubation period affects chitinase production; the enzyme activity rises to a maximum over a period of time, then drops with increasing time. The highest chitinase production was achieved after 7 days of incubation (3.56 ± 0.04 U/g). After this period, a reduction in the enzyme production due to metabolite degradation was observed [59]. The chitinase productivity was lost after the seventh day, and this may be attributed to the enzyme digestion by proteases, when the enzyme substrate in the culture medium was consumed [58].
The inoculation size (2.5 × 106 v/v) gave the highest production of chitinase about 3.94 ± 0.2 U/g. According to the findings of Shivalee et al. [60], there is a substantial rise in chitinase enzyme synthesis as the inoculum of Streptomyces pratensis increases. On the other hand, the lower inoculum size leads to fewer microbial cells in the production medium, which takes a longer time to proliferate and reach an optimal amount to consume the substrate and produce the desired output. In general, increasing the inoculum size promotes the growth- and growth-related activities of bacteria up to a certain point, after which there may be a drop-in microbial activity due to nutritional restriction [60].
The most pronounced production of chitinase was detected in media supplemented with yeast extract compared to the other additives. The addition of 100% cane and beet molasses gave the highest chitinase production by about 9.78 ± 0.4 and 11.56 ± 0.5 U/g, respectively. Beet molasses consist of a nitrogen content of 22.7, carbohydrates content of 12.87, Na of 328.5, K of 700.0, Ca of 17.76, Mg of 3.73, Fe of 1.09, Mn of 0.065, and Zn of 0.248 mg 100 mL−1. Our findings are consistent with those of Goksungur [61] who reported that the greatest chitosan extraction (783 ± 23 mg dm−3) from Rhizopus oryzae 00.4367 was recorded at the 72nd h of fermentation in molasses medium. This rises as a result of R. oryzae deamination of amino acids in molasses and the formation of ammonia, which raises the pH of the fermentation medium. Moreover, crab wastes are used in the simultaneous manufacture of chitin and L (+)-lactic acid via submerged fermentation of Lactobacillus sp. B2 with sugar cane molasses as a carbon source [62]. In addition, Sudhakar and Nagarajan [63] found that rice waste gave the highest production of chitinase from Serratia marcescens. The agricultural residue is cheap, plentiful, and easily accessible, and it provides superior nourishment to the microbes. Furthermore, the use of pure chitin raises the cost of enzyme manufacturing, which is a major barrier to the economic feasibility of lignocellulosic bioconversion and usage.
The maximum chitinase production was detected at 1.5 and 2.0% NaCl concentrations about 3.12 ± 0.1 and 3.88 ± 0.2 U/g, respectively, but the production decreased when NaCl increased or decreased above the optimum value. The chitinase production from T. funiculosus strain CBS 129594 in the presence of different NaCl levels exhibited good catalytic performance, proving its use in the successful management of chitinous biowastes. These results were in accordance with Halder et al. [55].
The chitinase from T. funiculosus strain CBS 129594 was effectively purified using 60% (NH4)2SO4, which resulted in the maximum protein recovery and chitinase recovered activity (73.3%), and 1.39-fold purification. The total protein and chitinase activity recovered from Sephadex G-100 achieved approximately 65.4% and 2.89-fold purification of the tested sample. The percent saturation of (NH4)2SO4 utilized for the purification of chitinase differs according to the isolates. Precipitation saturation ranged from 30% to 85% [64].
In-gel electrophoresis, the purified chitinase had a molecular mass of 45 kDa. This value fell within the range of molecular weights (35–74 kDa) of different fungal chitinases [65]. These results are in accordance with Chuan et al. [54], who used SDS-PAGE to purify two chitinases enzymes from T. flavus culture filtrate and reported that their molecular weights were 41 and 32 kDa, respectively.
The amino acid composition of the purified chitinase from T. funiculosus strain CBS 129594 revealed sixteen amino acids, with aspartic acid (15.20%) being the most abundant, followed by glutamic acid (12.3%), and methionine (1.6%) being the least abundant. These results are similar to Farag et al. [66] who observed that the pure chitinase synthesized by Aspergillus terreus contained 17 amino acids. Aspartic acid had the largest percentage, followed by glutamic acid.
T. funiculosus-purified chitinase had the highest activity at 40 °C (12.36 ± 0.5 U/mL), with a relative activity of 85.9% at 70 °C. The decline in chitinase activity at this temperature could be due to peptide chain hydrolysis, inaccurate confirmation, aggregation, or degradation of amino acids [66]. Our findings are completely consistent with the optimal temperature range for most fungal chitinase, which is 25 to 50 °C [52,67]. The type of the amino acids found in the active site determines pH dependent enzyme activity [68]. The results showed that pure chitinase was ideally active at pH 6.5, with an activity of 13.93 ± 0.9 U/mL and a relative activity of 90.6%. This optimum pH is the same as that recorded for purified chitinase from Thermomyces lanuginosus [69].
Metals have both a stimulating and an inhibiting influence on enzyme activity; therefore, metal compatibility profiling of an enzyme is required to determine optimal catalytic conditions. The purified chitinase was enhanced by Ca2+, Cu2+, Na+, Mn2+ and Mg2+ so it has acted as a potent activator. In addition, the purified chitinase activity was suppressed by Hg2+, Ag2+, and Li+, while Zn2+ and Co2+ did not cause any effect compared to media without any metals. These findings suggest that the chitinase under investigation is a metalloenzyme. These findings are consistent with those of Farag et al. [66] who discovered that various metal ions (Ca2+, Mn2+, Na+, K+, Mg2+, and Cu2+) can be used as activators for A. terreus pure chitinase. Other metals, on the other hand, decreased chitinase activity (Cd2+, Zn2+, pb2+, and Hg2+). Purified chitinase was stable and mildly more active by sodium dodecyl sulphate (SDS), Tween-80 as surfactants, EDTA as chelating agents, and PMSF as inhibitors. This could be attributed to the amino acid content as well as the three-dimensional structure, which supported the enzyme’s relative resistance to hydrophobic/non-polar microenvironments [68]. PMSF has a minor inhibition on chitinase activity suggesting that (1): serine or threonine hydroxyl groups are not implicated in substrate binding to the purified enzyme, and (2): a reducing agent such as iodoacetate shows evidence for the lack of -SH group in enzyme active sites [55].
The DPPH free radical is a stable free radical that is commonly used to estimate the free radical scavenging capabilities of antioxidants [34,70,71]. Chitin has exceptional antioxidative properties because it scavenges free radicals. This ability has been linked to the amino and hydroxyl groups, which can bind with unstable free radicals to produce stable macromolecular radicals [43]. The highest percent inhibition of DPPH and ABTS was obtained at a high concentration of purified enzymes (400 g/mL), which was approximately 57.8% and 63.7%, respectively. The IC50 value of DPPH radicals and ABTS of purified chitinase produced by T. funiculosus strain CBS 129594 was 199 and 306 μg/mL concentration, respectively.
Purified chitinase has antimicrobial activity against Gram-negative bacteria (P. aeruginosa, E. coli), Gram-positive bacteria (S. aureus, B. subtilis), and fungi (C. albicans, A. niger). Gram-negative bacteria are more sensitive than Gram-positive bacteria. These results are in accordance with those of Farag et al. [55], who found that pure chitinase synthesized from A. terreus exhibited a broad spectrum of antibacterial activity against S. aureus, S. typhi, and P. aeruginosa and antifungal activity against A. niger, A. oryzae, and Penicillum oxysporium. Furthermore, our findings are consistent with other chitinases obtained from multiple microbes [72]. Fungal chitinases have antifungal activity by inhibiting germ tube elongation, spore germination, hyphal tip formation, and spore rupture [73]. Halder et al. [43] explain that chitin is a crucial constituent of fungal cell walls, and chitinase is a well-known biocontrol agent that lyses the cell walls of pathogenic fungus in both live and dead conditions. Such deterioration can result in aberrant swellings of hyphae and spores, as well as mycelia lysis [43]. The most widely recognized mode of action of antibacterial activity is due to the free amino group and positive charge of chitin and chitosan-oligosaccharides, which can modify cell membrane permeability, causing the leaking of cell components and, ultimately, bacterial death [74]. The antibacterial properties of positively charged chitin and chitosan-oligosaccharides appear to be heavily influenced by the charge distribution of the bacterial cell wall. The bacterial cell wall is negatively charged. As a result, Gram-negative bacteria adsorb more positively charged chitin and chitosan-oligosaccharides on the surface due to their high negative charge than Gram-positive bacteria. This explains why chitin and chitosan-oligosaccharide combinations are more toxic to Gram-negative bacteria [75].
The highest concentrations of purified chitinase (1000 µg/mL) caused the higher toxicity of cancer cell line MCF7 (97%), HCT116 (88.2%), and HepG2 (97.1%). MTT assay showed that the purified chitinase from T. funiculosus strain CBS 129594 had a high toxic effect against breast cancer of the mammary gland (MCF7), hepatocellular cancer (HepG2), and a moderate cytotoxic activity against colorectal carcinoma colon cancer (HCT116). Similar results were obtained by Abu-Tahon and Isaac [76], who found that the purified chitinase from Trichoderma viride AUMC 13021 had an adverse effect on MCF7 with an IC50 value of 20 g/mL and HCT-116 cell lines with an IC50 value of 44 g/mL. This effect could be attributed to the presence of novel polycarbohydrates on the surface of cancer cells that interact with chitinase. Chitinase digests the carbohydrate portions of these glycoproteins or glycolipids, compromising their original function and causing tumor cells to die. The actual mechanism against cancer cell proliferation is unknown; however, it may be related to the electrostatic charges of chitosan-oligosaccharides, changes in tumor cell permeability, and modulation of tumor factor expression [77].

5. Conclusions

The chitinase enzyme was produced in solid-state fermentation using crustacean bio-wastes. The chitinase (45 kDa) was purified by simple procedures of precipitation, gel filtration, and ion exchange chromatography. The high concentration of purified enzymes (400 μg/mL) gave the highest % inhibition of DPPH and ABTS about 57.8% and 63.7% respectively. Moreover, the purified chitinase had antimicrobial activity against Gram-positive bacteria (P. aeruginosa, E. coli), Gram-positive bacteria (S. aureus, B. subtilis), and fungi (C. albicans, A. niger). Gram-negative bacteria are more sensitive than Gram-positive bacteria. In addition, the highest concentrations of purified chitinase (1000 µg/mL) caused higher toxicity against breast cancer of the mammary gland (MCF7), hepatocellular cancer (HepG2), and moderate cytotoxic activity against colorectal carcinoma colon cancer (HCT116). Therefore, we can state that chitinase is a promising candidate for antimicrobial activity, cancer therapy, and eco-friendly biocontrol agents, as an alternative to harmful chemical pesticides. To the best of our knowledge, this is the first report to determine the antioxidant, antimicrobial, and anticancer activity of purified chitinase from T. funiculosus strain CBS 129594.

Author Contributions

Conceptualization, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; methodology, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; software H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; validation, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; formal analysis, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; investigation, H.S.E.-B. and H.I.M., resources, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; data curation, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; writing—original draft preparation, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; writing—review and editing, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; visualization, H.S.E.-B., O.M.E.-M., H.I.M., and A.E.E.-A.; supervision, H.S.E.-B. and H.I.M.; project administration, H.S.E.-B.; funding acquisition, H.S.E.-B. All authors have read and agreed to the published version of the manuscript.

Funding

This work was supported through the annual funding track by the Deanship of Scientific Research, Vice Presidency for Graduate Studies and Scientific Research, King Faisal University, Saudi Arabia (Grant No. 1815).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data are contained within the article.

Acknowledgments

The authors thank the Deanship of Scientific Research, Vice Presidency for Graduate Studies and Scientific Research, King Faisal University, Saudi Arabia, for the financial support to conduct and publish this research. Also, the authors thank the Faculty of Education, Ain Shams University, and the Faculty of Agriculture, Cairo University for the financial support to conduct this research.

Conflicts of Interest

The authors declare no conflict of interest.

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Agronomy 12 02818 g001 550
Figure 1. Morphological (A,B) and molecular identification (C) of T. funiculosus strain CBS 129594.
Figure 1. Morphological (A,B) and molecular identification (C) of T. funiculosus strain CBS 129594.
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Figure 2. Effect of crab shellfish chitin and on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
Figure 2. Effect of crab shellfish chitin and on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
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Figure 3. Effect of the incubation period (A) and inoculum size (B) on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
Figure 3. Effect of the incubation period (A) and inoculum size (B) on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
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Figure 4. Effect of additives concentrations on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
Figure 4. Effect of additives concentrations on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
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Figure 5. Effect of sugar industry concentration on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
Figure 5. Effect of sugar industry concentration on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
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Figure 6. Effect of NaCl concentrations on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case-letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
Figure 6. Effect of NaCl concentrations on the production of chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD). Mean values in each bar followed by a different lower-case-letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
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Figure 7. SDS-PAGE of purified chitinase from T. funiculosus strain CBS 129594. Lane 1, molecular weight standards (Marker), Lane 2, purified chitinase.
Figure 7. SDS-PAGE of purified chitinase from T. funiculosus strain CBS 129594. Lane 1, molecular weight standards (Marker), Lane 2, purified chitinase.
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Figure 8. Amino acid composition of purified chitinase from T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD).
Figure 8. Amino acid composition of purified chitinase from T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD).
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Figure 9. Effect of temperature (A) and thermal stability (B) on the activity of purified chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD).
Figure 9. Effect of temperature (A) and thermal stability (B) on the activity of purified chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD).
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Figure 10. Effect of pH (A) and thermal stability (B) on the activity of purified chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD).
Figure 10. Effect of pH (A) and thermal stability (B) on the activity of purified chitinase on SSF by T. funiculosus strain CBS 129594. The values are the means of three replicates with standard deviation (±SD).
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Figure 11. Antimicrobial efficiency of purified T. funiculosus strain CBS 129594 chitinase against Pseudomonas aeruginosa (A), Escherichia coli (B), Bacillus subtilis (C), Staphylococcus aureus (D), Aspergillus niger (E), and Candida albicans (F).
Figure 11. Antimicrobial efficiency of purified T. funiculosus strain CBS 129594 chitinase against Pseudomonas aeruginosa (A), Escherichia coli (B), Bacillus subtilis (C), Staphylococcus aureus (D), Aspergillus niger (E), and Candida albicans (F).
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Figure 12. Effect of purified chitinase by T. funiculosus strain CBS 129594 on MCF7, HCT16, and HepG2 cells at different concentrations.
Figure 12. Effect of purified chitinase by T. funiculosus strain CBS 129594 on MCF7, HCT16, and HepG2 cells at different concentrations.
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Figure 13. (A). Effect of purified chitinase by T. funiculosus strain CBS 129594 on MCF7 cells at different concentrations. (B). Effect of purified chitinase by T. funiculosus strain CBS 129594 on HCT 116 cells at different concentrations. (C). Effect of purified chitinase by T. funiculosus strain CBS 129594 on HepG2 cells at different concentrations.
Figure 13. (A). Effect of purified chitinase by T. funiculosus strain CBS 129594 on MCF7 cells at different concentrations. (B). Effect of purified chitinase by T. funiculosus strain CBS 129594 on HCT 116 cells at different concentrations. (C). Effect of purified chitinase by T. funiculosus strain CBS 129594 on HepG2 cells at different concentrations.
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Table
Table 1. Effect of moisture content (MC) and shellfish waste as a substrate medium ratio on the production of chitinase on solid state fermentation technique by T. funiculosus strain CBS 129594.
Table 1. Effect of moisture content (MC) and shellfish waste as a substrate medium ratio on the production of chitinase on solid state fermentation technique by T. funiculosus strain CBS 129594.
SubstrateWeight of Substrate: Volume of Medium (w/v) *
1:11:21:31:41:5
** MC
(%)		Chitinase Activity (U/g Substrate)** MC
(%)		Chitinase Activity (U/g Substrate)** MC
(%)		Chitinase Activity (U/g Substrate)** MC
(%)		Chitinase Activity (U/g Substrate)** MC
(%)		Chitinase Activity (U/g Substrate)
Chitin (Sigma)55.451.7 ± 0.1a701.72 ± 0.2b77.142.35 ± 0.2a81.542.07 ± 0.1a84.521.51 ± 0.1a
Crab shell chitin56.361.3 ± 0.1b69.372.98 ± 0.2a76.661.56 ± 0.2b81.151.24 ± 0.1c84.191.02 ± 0.1b
Shrimp shell chitin56.360.17 ± 0.01c70.00.32 ± 0.05c77.140.46 ± 0.01c81.541.40 ± 0.1b84.520.92 ± 0.05b
Crayfish shell chitin55.450.04 ± 0.01d69.370.13 ± 0.02d76.660.26 ± 0.01d81.150.39 ± 0.05d84.190.31 ± 0.02c
The values are the means of three replicates with standard deviation (±SD). Mean values in each column followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05. * (w/v): Ratio weight of substrate: volume of optimized modified Czapek’s medium. ** (MC): Moisture content.
Table
Table 2. Purification of chitinase from T. funiculosus strain CBS 129594.
Table 2. Purification of chitinase from T. funiculosus strain CBS 129594.
Purification
Step		Protein
(mg/mL)		Activity
(U/mL)		Specific
Activity (U/mg Protein)		Recovery
%		Purification Fold
Cell Free Filtrate (CFF)4.14 ± 0.2a17.56 ± 0.6a4.23 ± 0.1d1001.0
Cell Free Dialysate (CFD)3.66 ± 0.3b16.23 ± 0.5b4.40 ± 0.1d92.41.04
Ammonium sulfate (60%)1.56 ± 0.1c12.86 ± 0.6c8.21 ± 0.2c73.31.93
Sephadex–G100 (S-G100)1.12 ± 0.1d9.66 ± 0.3d8.63 ± 0.2b65.42.89
DEAE cellulose1.00 ± 0.1d9.32 ± 0.3d9.32 ± 0.3a60.87.22
The values are the means of three replicates with standard deviation (±SD). Mean values in each column followed by a different lower-case letter are significantly different according to Duncan’s multiple range tests at p ≤ 0.05.
Table
Table 3. Effect of some metal ions and inhibitors on the activity of purified chitinase on solid state fermentation by T. funiculosus strain CBS 129594.
Table 3. Effect of some metal ions and inhibitors on the activity of purified chitinase on solid state fermentation by T. funiculosus strain CBS 129594.
Metal IonsConc. (mM)Relative Enzyme Activity (%)
Control0100 ± 5.0
Na+5120 ± 7.0
10110 ± 8.0
Mg2+5110 ± 6.0
1080.5 ± 5.0
Mn2+5115 ± 9.0
10105 ± 9.0
Hg2+570 ± 8.0
1045.5 ± 4.0
Zn2+5100 ± 5.0
10100 ± 5.0
Ca2+5150 ± 10.0
10120 ± 10.0
Co2+5100 ± 6.0
10100 ± 5.0
Cu2+5130 ± 8.0
1082.9 ± 6.0
Ag2+560 ± 4.0
1040 ± 4.0
Li+560 ± 3.0
1050 ± 4.0
EDTA592 ± 5.0
1090 ± 8.0
PMSF593 ± 6.0
1091 ± 5.0
SDS595 ± 7.0
1092 ± 9.0
Tween-80595 ± 7.0
1093 ± 8.0
The values are the means of three replicates with standard deviation (±SD). EDTA (chelator agent), PMSF (inhibitor).
Table
Table 4. Antioxidant activities of purified chitinase produced by T. funiculosus strain CBS 129594 as well as ascorbic acid against DPPH and ABTS at different concentrations.
Table 4. Antioxidant activities of purified chitinase produced by T. funiculosus strain CBS 129594 as well as ascorbic acid against DPPH and ABTS at different concentrations.
TreatmentConcentration
(µg/mL)		% Inhibition of
DPPH		IC50 (μg/mL)% Inhibition of ABTSIC50 (μg/mL)
Purified enzymes10048.73 ± 1.01998.87 ± 0.5306
20050.55 ± 2.029.77 ± 1.0
30055.07 ± 2.257.46 ± 1.5
40057.76 ± 1.963.73 ± 1.8
Ascorbic acid512.5 ± 1.030.325.2 ± 1.020.0
1028.9 ± 1.538.4 ± 1.2
2040.1 ± 1.648.5 ± 1.6
4060.2 ± 2.380.2 ± 1.9
Butylated hydroxytoluene (BHT)522.1 ± 0.919.035.3 ± 1.211.8
1035.3 ± 1.350.2 ± 1.4
2060.2 ± 1.865.9 ± 1.6
4080.5 ± 2.590.4 ± 2.2
The values are the means of three replicates with standard deviation (±SD).
Table
Table 5. Antimicrobial efficiency of purified chitinase by T. funiculosus strain CBS 129594 against Gram-positive bacteria, Gram-negative bacteria, and fungi.
Table 5. Antimicrobial efficiency of purified chitinase by T. funiculosus strain CBS 129594 against Gram-positive bacteria, Gram-negative bacteria, and fungi.
Pathogenic MicroorganismInhibitions Zones
(mm)		Con.
Gram-positive bacteria
Bacillus subtilis (ATCC 6633) 24 ± 1.317
Staphylococcus aureus v (ATCC 6538)20 ± 1.019
Gram-negative bacteria
Escherichia coli (ATCC 8739)29 ± 1.225
Pseudomonas aeruginosa (ATCC 90274)38 ± 1.522
Fungi
Candida albicans (ATCC 10221)25 ± 1.221
Aspergillus niger26 ± 1.416
The values are the means of three replicates with standard deviation (±SD). Antimicrobial activity was determined by using agar diffusion, disc diameter: 6.0 mm (100 µL was tested). 10 mg/mL of all samples was dissolved in normal saline (0.9% Na Cl) or DMSO. Control: Gentamycin.
Table
Table 6. Effect of purified chitinase by T. funiculosus strain CBS 129594 on MCF7, HCT116, and HepG2 cells.
Table 6. Effect of purified chitinase by T. funiculosus strain CBS 129594 on MCF7, HCT116, and HepG2 cells.
Cell LineConc. of Purified Chitinase µg/mLMean O.DST.EViability %Toxicity %IC50
µg/mL
Mcf7Control0.5530.0101000
10000.01670.0013.0197.0101.81
5000.01770.0033.2096.8
2500.0680.00812.3687.6
1250.1750.01031.6568.4
62.50.4590.01282.9417.1
31.250.5470.00698.921.08
HCT116Control0.3740.0081000
10000.0440.00411.7688.2411.45
5000.1250.00633.5166.5
2500.3040.00881.2818.7
1250.3660.00497.952.05
62.50.3720.00599.550.45
31.250.3740.004699.910.09
HepG2Control0.5990.00891000
10000.0170.00082.89297.1118.74
5000.020.0013.3496.7
2500.090.00515.7584.3
1250.2560.00442.7957.2
62.50.5730.00795.664.3
31.250.5980.00799.8390.167
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El-Beltagi, H.S.; El-Mahdy, O.M.; Mohamed, H.I.; El-Ansary, A.E. Antioxidants, Antimicrobial, and Anticancer Activities of Purified Chitinase of Talaromyces funiculosus Strain CBS 129594 Biosynthesized Using Crustacean Bio-Wastes. Agronomy 2022, 12, 2818. https://doi.org/10.3390/agronomy12112818

AMA Style

El-Beltagi HS, El-Mahdy OM, Mohamed HI, El-Ansary AE. Antioxidants, Antimicrobial, and Anticancer Activities of Purified Chitinase of Talaromyces funiculosus Strain CBS 129594 Biosynthesized Using Crustacean Bio-Wastes. Agronomy. 2022; 12(11):2818. https://doi.org/10.3390/agronomy12112818

Chicago/Turabian Style

El-Beltagi, Hossam S., Omima M. El-Mahdy, Heba I. Mohamed, and Abeer E. El-Ansary. 2022. "Antioxidants, Antimicrobial, and Anticancer Activities of Purified Chitinase of Talaromyces funiculosus Strain CBS 129594 Biosynthesized Using Crustacean Bio-Wastes" Agronomy 12, no. 11: 2818. https://doi.org/10.3390/agronomy12112818

APA Style

El-Beltagi, H. S., El-Mahdy, O. M., Mohamed, H. I., & El-Ansary, A. E. (2022). Antioxidants, Antimicrobial, and Anticancer Activities of Purified Chitinase of Talaromyces funiculosus Strain CBS 129594 Biosynthesized Using Crustacean Bio-Wastes. Agronomy, 12(11), 2818. https://doi.org/10.3390/agronomy12112818

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