|Year : 2019 | Volume
| Issue : 4 | Page : 320-331
Biosynthesis of biologically active chitinase utilizing some Egyptian chitinaceous wastes and the properties of the synthesized enzyme
Heba M Shalaby1, Salah A Abo-Sdera2, Saadia M Easa3, Abdel-Mohsen S Ismail1
1 Chemistry of Natural and Microbial Products Department, Pharmaceutical Industries Researches Division, National Research Centre, Cairo, Egypt
2 Agricultural Microbiology, National Rearsech Center, Cairo, Egypt
3 Microbiology Department, Faculty of Science, Ain Shams University, Cairo, Egypt
|Date of Submission||30-Apr-2019|
|Date of Acceptance||19-Jun-2019|
|Date of Web Publication||28-Jan-2020|
MSc Heba M Shalaby
Chemistry of Natural and Microbial Products Department, National Research Centre, Dokki, Giza 12622
Source of Support: None, Conflict of Interest: None
Background and objective Chitin-degrading enzymes have an utmost practical importance in many fields such as medicine, agriculture, and industry. These enzymes are used as effective antibacterial, antifungal, antihypertensive, and antioxidant agents and also as excellent food quality enhancers. The objective of the present article was to formulate the production medium and to pinpoint the proper growth conditions for the chosen microorganism producing highly active chitinase enzymes. The general properties of the crude enzyme preparation were determined to define the proper conditions for enzyme action. Under the specified conditions, the capability of the enzyme preparation for antimicrobial and antioxidant activities were decided.
Materials and methods Eighteen recommended microbial strains were screened for biologically active chitinolytic enzymes productivity. Chitinase enzyme was determined, and also the important properties of the crude chitinase were duly pinpointed. Finally, biological activities of the crude enzyme were studied.
Results and conclusion and conclusion Among all the 18 organisms, Streptomyces halstedii H2 was the most potent producer of an influential chitinase enzyme. The maximum chitinase activity of 49.5 U/ml was obtained from medium contains glucose 6 g/l, ammonium nitrate (0.9 g/l), and urea (0.64 g/l) at 30°C and pH 9.0. The important properties of the streptomycetal chitinase were duly pinpointed as follows: optimum enzyme and substrate concentrations were 1.6 mg/ml and 1.4% (w/v), respectively, and optimum reaction pH and temperature were 7.2 and 45°C, respectively. The crude preparation was stable for 60 min at pH 7.2 and 30°C and retained 92.6% of the original activity. Under the specified conditions, at varying concentrations, the enzyme preparation exhibited considerable 2, 2-diphenyl-1-picrylhydrazyl radical scavenging activity accompanied with low antimicrobial activity, pointing out the partial purification necessity of the crude enzyme preparation.
Keywords: biological activities, chitinase, 2, 2-diphenyl-1-picrylhydrazyl radical scavenging activity, marine chitinaceous wastes, microorganisms
|How to cite this article:|
Shalaby HM, Abo-Sdera SA, Easa SM, Ismail AMS. Biosynthesis of biologically active chitinase utilizing some Egyptian chitinaceous wastes and the properties of the synthesized enzyme. Egypt Pharmaceut J 2019;18:320-31
|How to cite this URL:|
Shalaby HM, Abo-Sdera SA, Easa SM, Ismail AMS. Biosynthesis of biologically active chitinase utilizing some Egyptian chitinaceous wastes and the properties of the synthesized enzyme. Egypt Pharmaceut J [serial online] 2019 [cited 2020 Aug 3];18:320-31. Available from: http://www.epj.eg.net/text.asp?2019/18/4/320/272270
| Introduction|| |
Marine wastes are considered the principal source of chitin, and more than 80 000 tons of chitin is produced from marine wastes every year . The accumulation of large amounts of wastes from crustaceans has become a main concern in the industry of seafood processing. Therefore, the utilization of these wastes for production of high value-added products, such as chitin, chitosan, and their derivatives for valuable applications in different fields, has vital demands ,,. In Egypt, crayfish, crabs, and shrimp are the most important crustacean seafoods. Shrimp shell wastes which constitute ∼40–50% of the total weight of shrimp become an environmental risk in Egypt owing to the recent increased consumption of shrimps ,,.
Chitin is nitrogen-containing polysaccharide consisting of β-1,4-linked N-acetyl-D-glucosamine, which is chemically analogous to the cellulose, except that one of the hydroxyl groups of each glucoside residue is replaced by an acetylated or deacetylated amino group. Chitin comprises ∼20–58% of the shellfish waste dry weight ,.
Chitinase (EC 220.127.116.11) enzymes are capable of hydrolyzing insoluble chitin to its oligo and monomeric components found in a variety of organisms including viruses, bacteria, fungi, insects, higher plants, animals, and play important physiological roles depending on their origin ,. Chitinases are constituents of several bacterial species; some of the best known genera include Aeromonas, Serratia, Vibrio, Streptomyces, and Bacillus ,,, and some fungal species ,. Chitinase applications in many fields, such as medical, agricultural, and industrial were comprehensively reported by many authors ,.
| Materials and methods|| |
The following microbes were screened in this study: Bacillus licheniformis, Bacillus megaterium, Bacillus subtilis, Pseudomonas putida, Pseudomonas fluorescens, Aspergillus niger, Aspergillus terreus, Dichotomous sp., Trichoderma harzianum, Trichoderma viride, Penicillium sp., Actinomadura sp., Nocardia sp., Streptomyces halstedii H2, Streptomyces natalensis NRC1, Streptomyces NRC 35, and Streptomyces sp., which were provided from the Culture Collection Centre of the National Research Centre, Egypt, whereas the bacterial strain B. subtilis NRRL 1315 was imported from the National Centre for Agricultural Utilization Research (Peoria, Illinois, USA; NRS).
On the other hand, the pathogenic bacterial strains (B. subtilis, Staphylococcus aureus, Escherichia coli, and Pseudomonas aeruginosa) and the pathogenic fungal strains (A. niger, Candida albicans, Fusarium oxysporum, and Rhizoctonia solani) were provided from Microbiology Department, Faculty of Science, Ain Shams University, Egypt.
Chitinaceous waste samples
Different chitinaceous wastes (shrimp, prawn, crab shells, and mollusk valves) were collected from local fish market in Cairo governorate, Egypt.
Chitinase production media
The following media were employed for the production of chitinases and were in the form of g/l.
Bacterial production medium 1
Colloidal chitin, 20; MgSO4.7H2O, 0.5; K2HPO4, 0.7; KH2PO4, 0.3; MnCl2, 0.001; ZnSO4, 0.001; and FeSO4.7H2O, 0.01, with pH of 7.2±0.2 .
Fungal production medium 2
Colloidal chitin, 10.0; yeast extract, 4.0; MgSO4.7H2O, 0.2; KH2PO4, 0.68; K2HPO4, 0.87; NH4Cl, 1.0; and KCl, 0.2, with pH of 6.5 .
Actinomycetal production media (3–6)
Chitin, 5.0; glucose, 3.0; soya bean, 3.9; NH4NO3, 0.9; MgSO4.7H2O, 0.3 and trace element solution, 1 ml; pH, 9.0. The trace element solution composed of (g/100ml): FeSO4.7H2O, 0.5; MnSO4.2H2O, 0.16; ZnSO4.7H2O, 0.14; and CaCl2, 0.2 .
Na-carboxy methyl cellulose, 10.0; peptone, 5.0; beef extract, 3.0; and NaCl, 5.0, with pH 7.2±0.2 .
Chitin, 10.0; KH2PO4, 1.36; yeast extract, 0.5; MgSO4.7H2O, 3.0; NaCl, 10.0; and (NH4) 2SO4, 1.0, with pH of 7.2±0.2 .
Chitin, 2.0; MgSO4.7H2O, 0.5; K2HPO4, 1.0; KCl, 0.5; (NH4)2SO4, 2.0; and FeSO4, 0.01, with pH of 7.2 .
The following two buffers were applied for different reaction pHs: phosphate buffer (pH 5.7–8.0) and glycine-NaOH buffer (pH 9.0–10.4).
Chitin and N-acetyl-D-glucosamine were purchased from MP Biomedical Inc, rue Geiler de Kayserberg 67402 illkirch, France and all the other chemicals were of analytical grade.
Different crustaceous wastes including shells and valves obtained from shrimps, prawns, crabs, and mollusks were washed under running water to remove organics, proteins, and other impurities. The specimens were then boiled in water for 0.5 h to remove the tissue, and followed by drying in an oven at 100°C for 2 h to break down the crystallinity of chitin. At the end, the dried shells were crushed in a blender with a sieve of 0.1 mm diameter. The raw material obtained is stored in bags and placed in a cold at 4°C to be used for microbial chitinase enzyme production .
Maintenance of the tested microorganisms and stock cultures
The tested bacteria were maintained on the nutrient agar slants and incubated at 37°C for 24 h, whereas the fungi were maintained on potato-dextrose-agar medium slants and incubated at 30°C for 7 days; other than that, the actinomycetes were maintained on yeast extract-malt extract medium (ISP2) slants and incubated at 30°C for 5 days.
Preparation of colloidal chitin
The colloidal chitin was prepared by the method described by Reid and Ogrydiazk  as follows: 5 g chitin was soaked in 100 ml of 85% (w/v) phosphoric acid at 4°C for 24 h. It was well mixed in an electric blender and diluted with distilled H2O, centrifuged at 2300 g for 20 min, washed several times with dist. H2O to remove of the excess acid, and then pH was adjusted to 7.0, using 1 N-NaOH. The colloidal chitin was dried at 45°C and kept in the refrigerator till use.
The protein of any solution was determined colourimetrically by Folin-Ciocalteu phenol reagent (Merck Company, Schuchardt, Germany) according to the method by Lowry et al. . The standard curve of protein was constructed using bovine serum albumin.
Chitinase activity assay
Chitinolytic enzyme activity was determined according to Stauff and Jaenicke  modified method, and the enzymatic products were measured applying the modified method of Neish , which was based on those described by Nelson  and Somogyi . This was done by incubating 1 ml of enzyme solution with 1 ml of (1%, w/v) colloidal chitin suspension in 0.2 M-phosphate buffer, at pH of 6.5. The reaction mixture was incubated in shaking water bath (70 rpm) at 40°C for 30 min, and then 1 ml of the mixture was withdrawn and added to 2 ml of copper reagent, boiled for 20 min, and cooled in running tap water, and then 2 ml of arsenomolybdate reagent was added and finally made up to 25 ml with distilled water. The released reducing sugars were colorimetrically measured at 520 nm. Standard curve was constructed using N-acetylglucosamine as standard. One chitinase activity unit was defined as 1 μmole N-acetylglucosamine released per one hour reaction at the specified conditions.
Optimization of enzyme production
Each of fermentation period, culture medium composition, initial pH, N and C sources, some saccharides addition, inoculum size and age, agitation rate, and incubation temperature were duly studied for microbial chitinase production optimization.
Employment of some pretreated chitinaceous marine wastes on microbial chitinase enzyme production
Equal weights of marine chitinaceous waste (shrimp shells, prawn shells, crab shell pieces, crab shells powder, or mollusk valves powder) were physically, chemically, and physicochemically pretreated, and then separately applied instead of the pure chitin in the optimized production medium.
General properties of the crude chitinase determination
Effects of enzyme protein and substrate (colloidal chitin) concentration, reaction temperature, and reaction pH on enzyme activity were studied, besides the enzyme pH stability.
Biological activities of the crude enzyme
Free radical scavenging activity
Free radical scavenging activity of the crude chitinase enzyme was determined by simple, rapid, and inexpensive method involving the use of the free radical, 2, 2-diphenyl-1-picrylhydrazyl (DPPH) with some modification . DPPH is a free radical of violet color. The antioxidants present in the sample scavenge the free radicals and turn it into yellow color. The color change from violet to yellow is proportional to the radical scavenging activity. In brief, the crude enzyme stock ethanolic solution (10.0 mg/ml) was diluted with ethanol to final concentrations ranged from 0.25 to 9.0 mg enzyme (wt/ml). Add 0.5 ml of a 0.3 mM-DPPH ethanolic solution separately to 0.5 ml of each sample solution, and the reaction mixture was vortexed and incubated for 1 h in room temperature. The absorbance of the solution was measured at 518 nm. Ascorbic acid was used as standard. The inhibitory percentage of DPPH was calculated according to the following equation:
where I═DPPH inhibition %, ABS CONTROL=absorbance of control (t=0 h), and ABS TEST=absorbance of a tested sample at the end of the reaction (t=1 h). Each assay was carried out in triplicate, and the results were averaged.
Percentage of radical scavenging activity was plotted against the corresponding concentration of the crude enzyme to obtain IC50 value. IC50 is the maximal concentration of the compound to cause 50% inhibition.
The antimicrobial activity of the tested crude S. halstedii H2 chitinase was carried out against gram-positive and gram-negative bacteria (B. subtilis, S. aureus and E. coli, and P. aeruginosa) and fungi (A. niger, C. albicans, F. oxysporum, and R. solani). Antimicrobial activity was determined by well diffusion method  with slight modification. Measurement of the inhibition zones was done by placing 0.1 ml of the crude preparation separately, of each one in 10-mm diameter wells cut in nutrient agar plates seeded by test bacteria and potato dextrose agar plates seeded by test fungi. After that, incubate all plates at 37°C for 24 h for bacteria and 30°C for 72 h for fungi, and the zone of inhibition was measured in mm diameter.
All experiments were repeated three times, the data shown in the corresponding tables and figures were the mean values of the experiments, the data statistics were analyzed and the Standard Error Mean (SEM) was determined.
| Results and discussion|| |
Screening for chitinase production by some recommended microorganisms
The screening of eighteen recommended microorganisms (six bacteria, six fungi, and six actinomycetes) for the production of chitinase enzymes in shaken cultures lasting for different periods (1–5, 3–7, and 1–6 days for bacteria, fungi, and actinomycetes, respectively), [Table 1],[Table 2],[Table 3], was performed. The data showed that with most cultures of different ages, the protein level was in the usual range and had no consistent relationship with both the microbial growth and chitinase activity. For all cultures, the final pH varied within a very limited range from the neutral to the slightly alkaline. It was noticed the prevalence of some tested actinomycetal followed by fungi to the bacterial organisms in chitinase enzymes production. Among the tested microorganisms, 5-day shaken S. halstedii H2 culture afforded the highest chitinase productivity (7.20 U/reaction) followed by the fungal species T. harzianum (4.12 U/reaction) on 3-day shaken culture. The other actinomycetes afforded in 2- and 3-day shaken cultures moderate chitinase productivity (2.26 U/reaction). In this respect, many streptomycetes were reported to be potent chitinase producers ,,. On the contrary, in a 3-L bench-top bioreactor, the fungal strain Penicillium janthinellum A8 produced highly active chitinase after 7 days .
|Table 1 Screening of some bacteria for chitinase activity during different incubation periods applying shaken culture technique|
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|Table 2 Screening of some fungi for chitinase activities during different incubation periods applying shaken culture technique|
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|Table 3 Screening of some actinomycetes for chitinase activity during different incubation periods applying shaken culture technique|
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Accordingly, among all the screened eighteen organisms, the S. halstedii H2 was chosen as the most potent chitinase producer in 5-day shaken culture and applied in the succeeding experiments.
Effect of fermentation period on enzyme production by Streptomyces halstedii H2
The highest chitinase activity and growth were achieved by S. halstedii H2 after 5 days of incubation (7.18 U/reaction) and gradually decreased at extended periods, where 6.4% 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 . This finding was more or less in accordance with those reported by Nawani and Kapadnis  and Suzuki et al. .
Effect of culture medium
Four culture media of varying formulations and including the foregoing basal medium 3 as control were examined to judge any was the most favorable and affording the highest chitinase productivity ([Figure 1]). The basal production medium 3 was the most favorable and led to the maximal S. halstedii H2 chitinase productivity (7.25 U/reaction). The other media 4 and 6 had almost lower similar effect on chitinase productivity, and more than 49% productivity decrease was recorded with medium 5. It was noticed that medium 4 enhanced S. halstedii H2 growth rather than the enzyme production. Accordingly, the culture medium 3 was chosen for chitinase production by S. halstedii H2. In this concern, similar formulation was designed by Suzuki et al.  for streptomycetal chitinase production by S. halstedii MF 425 applying the shaken technique. As medium 4 contained no chitin, the streptomycetal strain was able to produce considerable chitinase, indicating its constitutive nature, concluding that chitinase could be produced either adaptively or constitutively by S. halstedii H2.
|Figure 1 Effect of different culture media on Streptomyces halstedii H2 chitinase productivity applying 5-day-old shaken cultures; each value is presented as mean±SD (n=3).|
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Effect of initial pH
The effect of the culture medium initial pH on S. halstedii H2 chitinase productivity under wide pH range (5.7–10). The initial pH 9 afforded both the highest chitinase productivity and maximal dry weight of biomass (7.25 U/reaction and 750 mg/culture, respectively). The acidic pH or even slight acidic (5.7–6) had the most adverse effect on the enzyme production, and more than 67.6% loss in enzyme productivity was recorded. The optimum initial pH range 7–9 resulted in final alkaline pH range from 8.7 to 8.8. Conclusively, it seemed that the final alkalinity is inevitably needed for active chitinase production by S. halstedii H2. On the other hand, the changes in the protein value with the initial pH changes were limited. This was accorded with those reported by Mejía-Saulés et al. , who applied an initial pH from 6 to 9 for effective chitinase production by Serratia marcescens WF. In addition, initial pH of the production medium at 7–7.2 for optimal streptomycetal chitinase production was reported by Nawani and Kapadnis  and Suzuki et al. .
Effect of nitrogen source
Two groups of N sources on equal N basis were employed, that is, inorganic (ammonium nitrate, ammonium sulfate, and sodium nitrate) and organic (soya bean and urea). The data ([Figure 2]) disclosed that the replacement of ammonium nitrate in the basal medium 3 with ammonium sulfate or sodium nitrate led to 51.4 and 19.4% productivity loss. On the contrary, replacement of soya bean with urea led to noticeable jump in streptomycetal chitinase productivity, which was more than 4.7-fold as that of the basal medium. However, Nawani and Kapadnis  recommended both yeast extract and ammonium sulphate for streptomycetal chitinase production, and also Sherief et al.  decided the priority of peptone, yeast extract, and meat extract than urea for the fungal chitinase synthesis.
|Figure 2 Effect of N sources in the culture medium on Streptomyces halstedii H2 chitinase productivity.|
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Effect of carbon source
The data in [Figure 3] clarify that none of the added saccharides to the optimized medium favored S. halstedii H2 chitinase production as glucose did, except starch and lactose, which led to moderate enzyme yield. These findings were more or less coincided with those reported by many authors on the reverse effects of many carbon sources rather than chitin and chitin compounds, those including N-acetylglucosamine, glucose amine, glucose, galactose, mannose, and others on chitinase enzymes system of many microflora under controlled conditions ,. Nawani and Kapadnis  found 1% (w/v) chitin flakes from crab shells were proper for chitinase production by Streptomyces spp. NK528 and NK951, whereas Macagnan et al.  recommended the colloidal chitin for chitinase production from many streptomycetal species. Kuddus and Ahmed  found the same for Aeromonas hydrophila HS4 and Aeromonas punctata HS6 as the previous authors.
|Figure 3 Effect of C source in the culture medium on Streptomyces halstedii H2 chitinase productivity.|
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Effect of some additional saccharides
The effect of additional saccharides (glucose, lactose, and starch) in equal C basis in chitinase production medium was studied. Among the three sugars added, the highest chitinase productivity was obtained by the additional glucose (49.65 U/reaction) affording more than 44.75% productivity increase. In this respect, Kumar et al.  tested a mixture of different chitin forms (chitin and colloidal chitin) and found the ratio of 1 : 1.5 wt/wt, respectively, enhanced chitinase productivity by Humicola grisea ITCC.
Effects of inoculum size, age, and agitation rate
The effect of S. halstedii H2 inoculum size (5–15%, v/v) and age (12–72 h) was studied. 10% (v/v) inoculum (control experiment) was the most proper and led to the highest chitinase productivity, which decreased at above or below this concentration. Moreover, the inoculum age of 48 h was appropriate for the maximal chitinase productivity by S. halstedii H2 and the younger inoculum than 48 h produce lower enzyme yields. Suzuki et al.  inoculated S. halstedii culture medium with 4% (v/v) 47-h-old inoculum for optimal chitinase productivity. Concerning the effect of culture agitation rate, the rates from 150 to 250 rpm were applied in a bench-top thermostatic shaker, comparing it with a stationary culture, the shaking speed of 200 rpm resulted in the maximal chitinase activity, protein contents, well as dry weight of biomass, above which the productivity considerably decreased, and this was similar to those decided by EL-Masry .
Different incubation temperatures (25–50°C) effect on the streptomycetal chitinase productivity was followed up. The maximal productivity was achieved at 30°C. The same optimum incubation temperature (30°C) was reported in shaken cultures for other streptomycetal chitinase enzymes ,, whereas Sakai et al. , Yuli et al. , and EL-Masry  decided higher temperatures (58, 55, 50, and 40°C) for chitinase production by Bacillus sp. MH-1, Bacillus sp.13.26, Brevibacillus sp. RPSG-NRC KX350091, and Actinomadura sp., respectively.
Employment of physically and chemically pretreated marine chitinaceous wastes for streptomycetal chitinase production
The employment of some physically, chemically, and physicochemically pretreated marine chitinaceous wastes, namely, shrimp, prawn, crab shells (pieces and powder), and mollusk valves powder instead of chitin powder in the optimized fermentation culture medium was implemented in 0.5% (w/v) concentration as recommended by many authors ,. Physically pretreated (crushed and soaked overnight in distilled H2O) wastes were tested for the extracellular chitinase enzymes production by S. halstedii H2. The data ([Figure 4]a) declared that in all cases chitin powder was the most favorable by S. 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 alkali-treated previous wastes crushed and soaked overnight in 0.1 M-NaOH were also examined ([Figure 4]b); nevertheless, chitin powder was still the most favorable to S. halstedii H2 and led to the highest chitinolytic activity (49.65 U/reaction). Physically (crushed and H2O soaked overnight) or physicochemically (crushed and soaked in 0.1 M-NaOH overnight) pretreated crab shells to a great extent favored the streptomycetal chitinase synthesis as the pure chitin powder did ([Figure 4]a and b).
|Figure 4 Effect of some (a) physically pretreated and (b) alkali-treated marine chitinaceous wastes on Streptomyces halstedii H2 chitinase production.|
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In this concern, the utilization of the marine physically pretreated prawn and crab shells powder for microbial chitinase production was reported by many authors ,,. In addition, steam, drying, and milling were proceeded on prawn shells, crab shells, and squid pens to be utilized for the production of many value-added products including enzymes (chitinases, proteases, and nattokinase) through bioconversion means .
General properties of crude enzyme
The general properties of the crude streptomycetal chitinase include the effects of the enzyme protein concentration, substrate (colloidal chitin) concentration, the reaction pH, the reaction temperature, and pH stability. The results ([Figure 5]a) indicated that 100 µg enzyme protein in the reaction mixture was able to proceed the reaction yielding 3.36 U/ml after 30 min at 40°C and pH 6.5. However, 200 µg/reaction led to a noticeable jump in enzyme activity, which reached 2.3-fold as that afforded by the lower enzyme concentration. The enzyme activity parallelly increased with the enzyme concentration till 1.6 mg enzyme/reaction, which achieved the highest activity (69.8 U/ml). This pointed out that 1.6 mg enzyme protein/reaction was enough to consume all the substrate applied in the reaction.
|Figure 5 (a) Effect of enzyme protein concentration on the crude Streptomyces halstedii H2 chitinase activity. (b) Effect of colloidal chitin concn (%, w/v) on the crude Streptomyces halstedii H2 chitinase activity. (c) Effect of pH on the crude Streptomyces halstedii H2 chitinase activity. (d) Effect of reaction temperature on the crude Streptomyces halstedii H2 chitinase activity.|
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The effect of substrate (colloidal chitin) concentration on chitinase was illustrated in [Figure 5]b, where the substrate concentration enhanced the reaction rate according to the mass action law, to reach the maximal at 1.4% (w/v) colloidal chitin indicating the saturation of all the available enzyme active sites with chitin molecules. In this connection, Ghaly , Nampoothiri et al. , and Sandhya et al.  reported that the optimum colloidal chitin concentration for many streptomycetal and other microbial chitinases was from 0.5–1.5% (w/v).
The data in [Figure 5]c display the effect of reaction pH on the S. halstedii H2 chitinase at different pH values ranging from 5.7 to 10, applying 0.2 M-phosphate buffer pH (5.7–8.0) and 0.2 M-glycine-NaOH buffer pH (9.0, 10.0). The data proved that alkaline pH (7.2) was the most favorable for the enzyme performance, and above or below this pH resulted in a gradual loss in the enzyme activity. It is worthy to note that at extreme alkaline pH (10.0), the enzyme still retained 71.2% of the optimum (at 7.2), indicating the good alkalinity tolerance of the streptomycetal chitinase. In this respect, the optimum pH of different chitinases from five strains of Streptomyces ranged from 4 to 7 .
The data in [Figure 5]d disclose that heat till 45°C activates the enzyme reaction, and above which the enzyme activity slowly decreased. Elevation of temperature form 40°C (control) to 45°C led to more than 2.9% activation, indicating that 45°C was the optimum. At higher temperatures (55°C), the crude enzyme retained most its activity, which exceeds than 96.9% of the control (40°C). The crude enzyme retained also 83.8% at 65°C, indicating its high thermostability. The residual activity detected at temperatures higher than 65°C might be owing to the stabilizing effect of its substrate in the enzyme–substrate complex form . In this respect, temperature 40°C was reported for streptomycetal chitinases by El-Sayed et al.  from Streptomyces albovinaceus S-22 and by Gomes et al.  from Streptomyces sp. RC1071. In addition, temperature 50°C was reported for chitinases of some Streptomyces strains .
The pH stability study was carried out on the crude enzyme preparation of the optimized concentration (1.6 mg protein/reaction), where it was heated alone at 30°C and different pH values (5.7, 6.5, 7.2, and 8) applying 0.2 M-phosphate buffer for different incubation periods (30, 45, and 60 min). At the end of each incubation period, the pH of the enzyme solution was readjusted to the optimum reaction pH (7.2) by adding 0.1 M-NaOH or 0.1 M-HCl, and then used in the reaction as usual manner. The residual chitinase activity (%) was calculated compared with the normal enzymatic reaction applying unheated crude enzyme sample. The data in [Figure 6] clarify that the enzyme exhibited the highest stability and retained all its activity (100%) after 30 min heating at 30°C and pH 7.2. The enzyme at the aforementioned pH 7.2 was still highly stable and retained most of its original activity (92.6 and 92.6%) for 45 and 60 min, respectively. It is noteworthy that the enzyme still retained 90.5 and 95.7% of the original after 30 min heating at pH 6.5 and 8, respectively. Conclusively, these findings confirmed the enzyme excellent pH stability and indicated its superiority to other streptomycetal chitinases, which were thermostable at temperatures below 40°C ,. On the other hand, pH treatment of the enzyme at 5.7 was more harmful than that at the alkaline pH 8.0, so that the enzyme lost more than 61.2% of the original after 45 and 60 min at 30°C and pH 5.7, respectively and this was more or less accorded with those reported by Wang et al. , who reported that the chitinase CH1 from Pseudomonas sp. TKU015 had good stability at pH 5–7 and temperatures less than 50°C.
|Figure 6 pH stability of the crude Streptomyces halstedii H2 chitinase activity.|
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2, 2-diphenyl-1-picrylhydrazyl radical scavenging activity
The DPPH free radical is a stable free radical, which has been widely accepted as a tool for estimating the free radical scavenging activities of antioxidants. The data in [Figure 7] disclose the DPPH radical scavenging activity of the crude chitinase enzyme, which ranged from 14.43 to 94.55% at varying concentrations from 0.25 to 10 mg enzyme preparation wt/ml in ethanol solution, whereas ascorbic acid displayed more than 90% at the concentration 5 mg/ml. The DPPH radical scavenging activity increased in an enzyme dose-dependent manner. In this respect, the antioxidant activity of Alcaligenes faecalis AU02 culture supernatant was determined through scavenging ability on 1,1-diphenyl-2-picrylhydrazyl (DPPH) as 84%, and the antioxidant compound was characterized by TLC .
|Figure 7 2, 2-diphenyl-1-picrylhydrazyl radical scavenging activity of the crude chitinase enzyme and ascorbic acid.|
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According to [Figure 7], the IC50 value was calculated for the applied crude chitinase to be at 0.95 mg, which contains 160 μg enzyme protein. It could be decided here that the S. halstedii H2 chitinase in its crude form exhibited a considerable scavenging activity comparing with the other related studies .
Concerning the antifungal activity, unfortunately the crude enzyme afforded low activity, and this will be followed up with the extended studies on the partially purified enzyme form, which is expected to exhibit good antimicrobial activity.
This work supported by Chemistry of Natural and Microbial Products Department, NRC, Egypt and Microbiology Department, Faculty of Science, Ain Shams University, Egypt. This work also was financially supported by Science and Technology Development Fund (STDF), of the Academy of Scientific Research and Technology, Egypt (Grant No. 6557).
Financial support and sponsorship
Conflicts of interest
There are no conflicts of interest.
| References|| |
Rinaudo M. Chitin and chitosan: properties and applications. Prog Polym Sci 2006; 31:603–632.
Shahidi F, Arachchi JKV, Jeon YJ. Food applications of chitin and chitosans. Trends Food Sci Technol 1999; 10:37–51.
Shahidi F, Abuzaytoun R. Chitin, chitosan and co-products: chemistry, production, applications and health effects. Adv Food Nutr Res 2005; 49:93–135.
Kim SK, Venkatesan J. Introduction to seafood processing by-products. In: Kim SK, ed. Seafood processing by-products: trends and applications ISBN: 9781461495901. New York, USA: Springer 2014. 1–9
Shahidi F, Synowiecki J, Naczk M. Utilization of shellfish processing discards. In: Bligh EG, Ackman RG eds. Seafood science and technology ISBN: 9780852381731. Oxford, UK: Fishing News Books 1992. 300–304
Sadek S, Rafael R, Shakouri M, Rafomanana G, Ribeiro FL, Clay J. Shrimp aquaculture in Africa and the Middle East: the current reality and trends for the future. Report prepared under the World Bank, NACA, WWF and FAO Consortium Program on Shrimp Farming and the Environment 2002; 1–42
Gallert Xu YC, Winter J. Chitin purification from shrimp wastes by microbial deproteination and decalcification. Appl Microbiol Biotechnol 2008; 79:687–697.
Wang SL, Chang WT. Purification and characterization of two bifunctional chitinases/lysozymes extracellularly produced by Pseudomonas aeruginosa K-187 in a shrimp and crab shell powder medium. Appl Environ Microbiol 1997; 63:380–386.
Saima MK, Roohi IZA. Isolation of novel chitinolytic bacteria and production optimization of extracellular chitinase. J Genetic Eng Biotechnol 2013; 11:39–46.
Goodday GW. The ecology of chitin degratiotic. in: Marshall KC ed. Advance in microbiol ecology. New York: New Plenum Press 1990. 11: 387–430.
Duo-Chuan L. Review of fungal chitinases. Mycopathologia 2006; 161:345–360.
Gohel V, Chaudhary T, Vyas P, Chatpar HS. Isolation and identification of marine chitinolytic bacteria and their potential in antifungal biocontrol. Indian J Exp Biol 2004; 42:715–720.
Wang SL, Lin TY, Yen YH, Guo CT, Chienyan H. Production of antifungal materials by bioconversion of shellfish chitin wastes fermented by Pseudomonas fluorescens
K-188. Enzyme Microb Technol 2005; 36:49–56.
Wang SL, Lin TY, Yen YH, Liao HF, Chen YJ. Bioconversion of shellfish chitin waste for the production of Bacillus subtilis
W-188 chitinase. Carbohydr Res 2006; 341:2507–2515.
Al-Ahmadi KJ, Yazdi MT, Najafi MF, Shahverdi AR. Optimization of medium and cultivation conditions for chitinase production by the newly isolated Aeromonas
sp. Biotechnology 2008; 7:266–272.
Patidar P, Agrawal D, Banerjee T, Patil S. Optimisation of process parameters for chitinase production by soil isolates of Penicillium chrysogenum
under solid substrate fermentation. Process Biochem 2005; 40:2962–2967.
Rattanakit N, Yano S, Plikomol A, Wakayama M, Tachiki T. Purification of Aspergillus
sp.S1-13 chitinases and their role in saccharification of chitin in mash of solid-state culture with shellfish waste. Biosc Bioeng 2007; 103:535–541.
Bhattacharya D, Nagpure A, Gupta RK. Bacterial chitinases: properties and potential. Crit Rev Biotechnol 2007; 27:21–28.
Russell G. Review of the application of chitin and its derivatives in agriculture to modify plant-microbial interactions and improve crop yields. Internet J Agronomy 2013; 3:757–793.
Murthy N, Bleakley B. Simple method of preparing colloidal chitin used for screening of chitinase producing microorganisms. Internet J Microbiol 2012; 10:1–4.
Zhang M, Puri AK, Govender A, Wang Z, Singh S, Permaul K. The multi-chitinolytic enzyme system of the compost-dwelling thermophilic fungus Thermomyces lanuginosus
. Proc Biochem 2015; 50:237–244.
Shalaby HM. Microbial utilization of some environmental chitinaceous wastes for the production of valuable products [MSc Thesis]. Al-Azhar, Egypt: Faculty of Science, Al-Azhar University; 2011.
Narasimhan A, Shivakumar S. Optimization of chitinase produced by a biocontrol strain of Bacillus subtilis
using Plackett-Burman design. Eur J Exp Biol 2012; 2:816–865.
Singh AK, Mehta G, Chhatpar HS. Optimization of medium constituents for improved chitinase production by Paenibacillus
sp D1 using statistical approach. Soc Appl. Microbiol 2009; 49:708–714.
Suzuki S, Nakanishi E, Furihata K, Miyamoto K, Tsujibo H, Watanabe T et al.
Chitinase inhibitor allosamidin promotes chitinase production of Streptomyces
generally. Int J Biol Macromol 2007; 43:13–19.
Mukherjee DP. Method for producing chitin or chitosan. United State Patent 2002; 6:310.
Reid JD, Ogrydiazk DM. Chitinase-over producing mutant of Serratia marcescens
. App Environm Microbial 1981; 41:644–669.
Lowry OH, Rosebrough NJ, Farr AL, Randall RJ. Protein measurement with the Folin phenol reagent. J Biol Chem 1951; 193:265–275.
Stauff J, Jaenicke R. Biochemisches Taschenbuch. Physikalische Chemie der Lösungen Berlin Heidelberg New York 1964; 2:93–102.
Neish AC. Analytical methods for bacterial fermentations. Nat Res Counc Canada Rep 1952; 46:3–8.
Nelson N. A photometric adaptation of the Somogyi method for the determination of glucose. J Biol Chem 1944; 153:375–380.
Somogyi MJ. Determination of blood sugar. J Biol Chem 1945; 16:61–63.
Peng C, Chen S, Lin F, Lin Z. Detection of antioxidative capacity in plants by scavenging organic free radical DPPH. Prog Biochem Biophys 2000; 27:57–61.
Perez C, Anesini C. In vitro antimicrobial activity of Argentine folk medicinal plants against Salmonella typhi
. J Ethno pharmacol 1994; 44:41–46.
Nawani NN, Kapadnis BP. Optimization of chitinase production using statistics based experimental designs. Process Biochem 2004; 40:651–660.
Mukherjee G, Sen SK. Purification, characterization, and antifungal activity of chitinase from Streptomyces venezuelae
. Curr Microbiol 2006; 53:265–269.
Fenice M, Leuba J, Federici F. Chitinolytic enzyme activity of Penicillium janthinellum
P9 in bench-top bioreactor. J Ferment Bioeng 1998; 86:620–623.
Ferdinand W. The enzyme molecule. London, New York: Wiley J and Sons Company. 1976; 224–227.
Mejía-Saulés J, Waliszwski KN, Garcia MA, Cruz-Camarillo R. The use of crude shrimp shell powder for chitinase production by Serratia marcescens
WF. Food Technol Biotechnol 2006; 44:95–100.
Sherief AA, El-Sawah MM, Abdel-Naby MA. Some properties of chitinase produce by a potent Aspergillus carneus
strain. Appl Microbiol Biotechnol 1991; 35:228–230.
Escott GM, Hearn VM, Adams DJ. Inducible chitinolytic system of Aspergillus fumigatus
. Microbiology 1998; 144:1575–1581.
EL-Masry HMM. Eco-friendly microbial biosynthesis of chitinases and formation of [N-acetylglucoseamine] [MSc thesis]. Ismailia, Egypt: Faculty Science, Suez Canal University; 2017.
Macagnan D, Romeiro RDS, Pomella AW. Production of lytic enzymes and siderophores and inhibition of germination of basidiospores of Moniliophthora
by phylloplane actinomycetes. Biol Control 2008; 47:309–314.
Kuddus SM, Ahmad RIZ. Isolation of novel chitinolytic bacteria and production optimization of extracellular chitinase. J Genetic Eng Biotechnol 2013; 11:39–46.
Kumar M, Brar A, Vivekanand V, Pareek N. Production of chitinase from thermophilic Humicola grisea
and its application in production of bioactive chitooligosaccharides. J Biol Macromol 2017; 104:1641–1647.
Sakai K, Yokota A, Kurokawa H, Wakayama M, Moriguchi M. Purification and characterization of three thermostable endochitinases of a noble Bacillus
strain MH-1, isolated from chitin-containing compost. Appl Env Microbiol 1998; 64:3397–3402.
Yuli PE, Suhartono MT, Rukayadi Y, Hwang JK, Pyun YR. Characteristics of thermostable chitinase enzymes from the Indonesian Bacillus
sp. 13.26. Enz Microb Technol 2004; 35:147–153.
Tarafdar A, Biswas G. Extraction of chitosan from prawn shell wastes and examination of its viable commercial applications. Int J Theoret Appl Res Mech Eng 2013; 2:17–24.
Kim SK, Venkatesan J. Introduction to seafood processing by products. In: Kim SK, ed. Seafood processing by-products: trends and applications ISBN: 9781461495901. New York, USA: Springer. 2014. 1–9
Chang WT, Chen CS, Wang SL. An antifungal chitinase produced by Bacillus cereus
with shrimp and crab shell powder as carbon source. Curr Microbiol 2003; 47:102–108.
Wang J, Zhang J, Song F, Gui T, Xiang J. Purification and characterization of chitinases from ridgetail white prawn Exopalaemon carinicauda
. Molecules 2015; 20:1955–1967.
Wang SY, Moyne AL, Thottappilly G, Wu SJ, Locy RD, Singh NK. Purification and characterization of a Bacillus cereus
exochitinase. Enz Microb Technol 2001; 28:492–498.
Ghaly MF. Improvement of the activity of cell wall degrading chitinase produced by Streptomyces albovenaceus
and its uses as antifungal and insecticide. N Egypt J Microbiol 2003; 7:190–197.
Nampoothiri KM, Baiju TV, Sandhya C, Sabu A, Szakacs G, Pandey A. Process optimization for antifungal chitinase production by Trichoderma harzianum
. Process Biochem 2004; 39:1583–1590.
Sandhya C, Binod P, Nampoothiri KM, Szakacs G, Pandey A. Microbial synthesis of chitinase in solid culture and its potential as biocontrol agent against phytopathogenic fungus Colletotric humgloeo sporioides
. App Biochem Biotechnol 2005; 127:1–15.
Hoster F, Schmitz J, Daniel R. Enrichment of chitinolytic microorganisms: isolation and characterization of a chitinase exhibiting antifungal activity against phytopathogenic fungi from a novel Streptomyces
strain. Appl Microbiol Biotechnol 2005; 66:434–442.
El-Sayed ES, Ezzat SM, Ghaly MF, Mansour M. Purification and characterization of two chitinases from Streptomyces albovinaceus S-22. World J of Microbiol Biotechnol 2000; 16:87–89.
Gomes RC, Semedo LTAS, Soares RMA, Linhares LF, Ulhoa CJ, Alviano CS, Coelho RRR. Purification of a thermostable endochitinase from Streptomyces
RC1071 isolated from a cerrado soil and its antagonism against phytopathogenic fungi. J Appl Microbiol 2001; 90:653–661.
El-Sayed SA, Ezzat SM, Ghaly MF, Mansour M, El-Bohey MA. Purification and characterization of two chitinases from Streptomyces albovinaceus
S-22. J Union Arab Biol 1999; 8:149–163.
Kim KJ, Yang YJ, Kim JG. Purification and characterization of chitinase from Streptomyces
sp.M-20. J Biochem Mol Biol 2003; 36:185–189.
Wang SL, Chen SJ, Wang CL. Purification and characterization of chitinases and chitosanases from a new species strain Pseudomonas
sp. TKU015 using shrimp shells as a substrate. Carbohydrate Res 2008; 343:1171–1179.
Neelamegam A, Mayavan VR, Shanmugam V, Thangavel B. Purification and characterization of chitinase from Alcaligenes faecalis AU02 by utilizing marine wastes and its antioxidant activity. Ann Microbiol 2011; 61:801–807.
[Figure 1], [Figure 2], [Figure 3], [Figure 4], [Figure 5], [Figure 6], [Figure 7]
[Table 1], [Table 2], [Table 3]