A novel bioactive edible coating based on sodium alginate and galbanum gum incorporated with essential oil of Ziziphora persica: The antioxidant and antimicrobial activity, and application in food model.
Galbanum, an aromatic gum resin, is produced from umbelliferous Persian plant species in the genus Ferula with the common Persian name “Barije”, chiefly endemic flora of the mountain ranges of northern Iran. Ziziphora persica is an edible medicinal plant, which is widely distributed in Iran, Turkey and Eurasian countries. The aims of the present research were to produce novel bioactive edible coatings based on sodium alginate (Alg), galbanum oleo-resin gum (GG) and the biocomposite of alginate and galbanum (CAG) containing different concentration of Ziziphora essential oil (ZEO), and evaluate the antioxidant and the antimicrobial activities of these substances in vitro and their effect on the quality and shelf life of chicken fillet during cold storage. Overall, the MIC and MBC values of GG and ZEO extracts ranging from 1.25 to 50 (µg mL-1) proved that Gram-positive bacteria were more susceptible to GG and ZEO than Gram-negative bacteria. The high phenolic and flavonoid contents and antioxidant activities of GG and ZEO were proved by in vitro analysis. Results showed that coatings of Alg had no significant effect on decreasing the microbial load of aerobic mesophilic and psychrotrophic bacteria, lactic acid bacteria, Pseudomonas spp., and Enterobacteriaceae, as well as L.monocytogenes (p>0.05), while the coating of fillet with GG, CAG alone and in combination with ZEO showed a significant differences with the other treatments during 12 days of storage (p<0.05). The results also showed that TVBN, TBARS and peroxide formation in the samples treated by GG/ZEO was significantly lower than other groups (p<0.05).
Keywords: Ferula gummosa galbanum gum; Sodium alginate; Ziziphora persica essential oil; Antioxidant and antimicrobial activities; Shelf life of chicken fillet
In addition to the extensive research to identify and replace natural and plant-based with chemical preservatives, edible coatings and biodegradable films based on proteins and polysaccharides have recently gained more interest in food preservation due to their antimicrobial and antioxidant activities (Cazón, Velazquez, Ramírez, & Vázquez, 2017). The usage of several bio-polysaccharides in food preservation has been studied, including alginate, carrageenan, cellulose, chitosan, kefiran, pectin, pullulan and starch (Cazón et al., 2017). Alginates are the salts of alginic acid, a linear co-polymer composed of 1-4β-d-mannuronic acid (M) and α-l-guluronic acid (G) that refined from brown seaweeds called Phaeophyceae, by treatment with aqueous alkali solutions, typically with NaOH (Lee & Mooney, 2012). In addition to the algal origin, alginate can be produced by Azotobacter and Pseudomonas with tailor-made features and wide applications in biomedical applications (Maleki, Almaas, Zotchev, Valla, & Ertesvåg, 2016; Trujillo-Roldán, Monsalve-Gil, Cuesta-Álvarez, & Valdez-Cruz, 2015).
Alginate absorbs water quickly and used as an ingredient in a variety of manufacturing industries like as paper, textiles, pharmaceutical, dentistry, prosthetics, lifecasting and different types of medical products including wound healing, drug delivery, and tissue engineering materials (Cui et al., 2016; Lee & Mooney, 2012; Meng et al., 2017; Rehim, El-Samahy, Badawy, & Mohram, 2016). In the food industries, sodium alginate used as a thickening agent for drinks, ice creams, cosmetics, sauces, salads, puddings, jams, tomato sauces and canned products, and as a gelling agent for jellies. Alginate gel can be made into a variety of foods, maintain a good colloidal form, and suitable for frozen and man-made foods (Draget, Moe, Skjak-Bræk, & Smidsrød, 2016).
The term active packaging usually means the inclusion of specific compounds with active function beyond the packaging materials to improve safety and quality, as well as to extend the shelf life of the foodstuffs (Fang, Zhao, Warner, & Johnson, 2017). The use of antimicrobials in food packaging has more advantages compared to direct adding of these agents to food products because the antimicrobial agents added to surfaces of food products by sprays or drops are not effective enough to inhibit microorganisms (Sung et al., 2013). This is attributable to the rapid spread of the antimicrobial agents into foods and denaturation of the active compounds by food components which reduce the reactivity of the functional agents. Furthermore, antimicrobial packaging provides slow and continuous migration of these agents from packaging materials to food surfaces capable of maintaining an antimicrobial agent at high concentrations over a long period of time (Quintavalla & Vicini, 2002). On the other hand, natural antimicrobial ingredients like essential oils (EOs) are volatile components and can not be used alone. For enhancing the efficacy of these materials using active films such as bio-polysaccharides is recommended (Sánchez-Ortega et al., 2014).
Galbanum, an aromatic gum resin, is produced from umbelliferous Persian plant species in the genus Ferula with the common Persian name “Barije”, chiefly endemic flora such as Ferula gummosa, Ferula persica, and Ferula tabasensis that grow extensively on the slopes of the mountain ranges of northern Iran (Asili, Sahebkar, Bazzaz, Sharifi, & Iranshahi, 2009). Galbanum resin has a very bitter, acrid, and peculiar scent followed by a very intense green, spicy, woody, balsam-like fragrance. Galbanum was used traditionally as antiseptic, anti-flatulent, anti-seizure agent, anti-spasm, pain-killer, inflammation reliever, and tonic of memory enhancement by the ancient Iranians and Egyptians (Mahboubi, 2016). In more recent studies, the antimicrobial, anti-inflammatory, anti-convulsant, anti-nociceptive, anti-leptic, anti-septic, analgesic, and anti-diabetic, spasmolytic, carminative, expectorant, anti-catarrh, anti-rheumatic, anti-nociceptive, anti-hysteric, laxative, aphrodisiac and many other medicinal applications of galbanum have been reported (Abbaszadegan et al., 2015; Adhami et al., 2014; Enauyatifard, Azadbakht, & Fadakar, 2012; Gudarzi et al., 2015; Moosavi et al., 2015). Galbanum is also used in the manufacture of textiles, cosmetics, and various glues (Mortazaienezhad & Sadeghian, 2006).
The genus Ziziphora (Lamiaceae family) with the common Persian name “kakuti-e kuhi” consists of four species (Z. clinopodioides, Z. capitata, Z. persica and Z. tenuior). Ziziphora persica is an annual herb and edible medicinal plant, which is widely distributed in Iran, Turkey and Eurasian countries. It is known as a medical plant that its leaves, flowers and stems frequently used as wild vegetable or additive in foods offer aroma and flavor (Dakah, Zaid, Suleiman, Abbas, & Wink, 2014). A number of Ziziphora species have been used for thousands of years in the traditional system of medicine as infusions for various purposes such as sedative, stomachic tonic, heart disorders, common cold, diarrhea, coughing, antiseptic, wound healing and carminative among others (Khodaverdi-Samani, Pirbalouti, Shirmardi, & Malekpoor, 2015). The chemical composition, antibacterial, antioxidant and antifungal activities of the essential oil of Ziziphora species (endemic Iranian herbs) has already been studied (Ebrahimi, Hadian, & Sonboli, 2009; Ganjali, Harati, & Kaykhaii, 2016; Khodaverdi-Samani et al., 2015; Shahbazi, 2016). Therefore, the objectives of the present research were to produce bioactive edible coating based on alginate (Alg), galbanum oleo-resin gum (GG) and the biocomposite of alginate and galbanum (CAG) containing different concentration of Ziziphora persica essential oil (ZEO), and evaluate the antioxidant and the antimicrobial activities of these substances in vitro and their effect on the quality and shelf life of chicken fillet during cold storage at 4˚C.
2. Materials and methods
2.1. Materials and chemicals
Galbanum (Ferula gummosa) was prepared from Mazandaran province, Iran. The plant Ziziphora persica (aerial parts) was collected at the flowering stage from Jolfa city (West Azerbaijan Province, Iran) during May-June 2016. The gum resin and plant materials were further identified at the Pharmacognosy Department, Faculty of Pharmacy, Tehran University of Medical Sciences, Tehran, Iran. Sodium alginate was purchased from Sigma-Aldrich Chemical Co. (Oakville, ON, USA). All other chemicals used were of analytical grade and were provided by Merck Chemical Co. (Darmstadt, Germany).
2.2. Extraction of gum from galbanum
The extraction of oleo-gum-resin from F. gummosa was performed according to the method described by Jalali et al. (2013). The exudates were collected by scratching the surface of the plant near the root and extracted with diethyl ether solvent. The samples were dried by a rotary evaporator and the powdered dried galbanum was macerated in distilled water at 50˚C and shaken for 30 min. Afterward, the mixture was stirred for 24 h and filtered through Whatman # 4 filter paper. The filtrate gum was stored at 4°C until analysis.
2.3. Chemical analysis of gum
The Moisture, ash, crude protein and fat contents of the oleo-gum-resin were determined according to AOAC (2005). Monosaccharide determination was performed after hydrolyzing the gum by acids (0.5 M sulphuric acid, 80 °C, 18 h). The hydrolysates were neutralized with NaOH solution and then centrifuged at 1500g for 10 min and the supernatant was filtered through a 0.45 μm filter membrane. The composition of the monosaccharides was determined by HPLC system with an RI detector (Knauer, Germany) equipped with a Eurokat H carbohydrate analysis column (300×8 mm, Knauer, Germany). The mobile phase was acidified water (0.01N sulfuric acid, pH 2) at a flow rate of 0.4 mL min-1 with a column temperature of 75°C.
2.4. Extraction and chemical analysis of Ziziphora essential oil
The aerial parts of the plant with 650 mL water were placed in round flask and the essential oil was extracted through hydro-distillation by using a Clevenger-type apparatus 3.5 h and continued to produce oil in a yield of 1.05% (v/v). The oily layer obtained on top of the aqueous distillate was separated and anhydrous sodium sulfate was used to dry the obtained essential oil and stored at 4°C until analysis. The oily layer obtained on top of the aqueous distillate was separated and dried over anhydrous Na2SO4 and stored in sealed vials under refrigeration prior to analysis.
Gas chromatography (GC) analyses were performed on Ziziphora persica essential oil samples by using a Shimadzu QP-2010 GC-MS system equipped with DB-5 ms capillary column (length, 30 m; column i.d., 0.25 mm; film thickness, 0.25 µm) and connected to a flame ionization detector. The temperature of both injector and detector was 250°C. High purity helium was served as the carrier gas (1.0 mL min-1). The column temperature was kept 40°C for 5 min, then heated to 250°C at a rate of 5°C min-1 and finally kept constant at 250 °C for 5 min. The split ratio was 1:100. GC-MS spectra were taken at 70 eV (E1) over the mass range 30-350 amu using an electron multiplier voltage of 1800 eV and scan time was 2 scans per second. For each of the substances analyzed, three characteristic ions were monitored during the analysis (Ding, Yang, Liu, & Tian, 2014).
2.5. Determination of the MIC and MBC
The minimum inhibitory concentration (MIC) and minimum bactericide concentration (MBC) of galbanum gum (GG) and Ziziphora essential oil (ZEO) were measured by the broth microdilution method using 96-well microtiter plates as described by the Clinical and Laboratory Standards Institute (CLSI, 2012). The GG and ZEO were dissolved in Mueller-Hinton broth (MHB) supplemented with Tween 80 at a final concentration of 0.1% (v/v). Dilutions of the samples (0.1–50%) were prepared in test tubes and dispensed into the wells of a microtiter plate according to a checkerboard design; each well was then inoculated with 100 mL of the bacterial suspensions include; 3 Gram-negative (E. coli, P. aeruginosa, and S. Typhimurium) and 3 Gram-positive bacteria (L. monocytogenes, B. cereus, and S. aureus) with ca. 5×105 CFU mL-1 concentration. After incubation at 35°C for 24 h, the wells were examined by microplate reader (Bioteck Instruments, Inc. VT, USA) for the growth of microorganism (absorbance at 550 nm) and the MICs were determined. The MIC was defined as the lowest concentration at which the microorganism did not demonstrate visible growth. MBCs were determined by plating 10 µL from each well demonstrating no visible growth. The MBC was defined as the lowest concentration of antimicrobials that killed at least 99.9% of the initial inoculums. Each experiment was repeated three times.
2.6. Determination of total phenolic and flavonoid contents
The total phenolic contents of GG and ZEO were determined by the Folin–Ciocalteu reagent with gallic acid as a standard (Marinova, Ribarova, & Atanassova, 2005). Briefly, 0.1 ml of each extract (0.1 mg ml-1) or standard gallic acid solutions (0–0.5 mg) were mixed with 1 ml Folin–Ciocalteu reagent (10%) for 5 min and 0.3 ml Na2CO3 (10%) was then added. The absorbance of the reaction was measured at 765 nm after 2 h of incubation at room temperature by using a UV-VIS spectrophotometer (2550, Shimadzu, Japan). Total phenolic content was expressed as mg of gallic acid equivalent per gram extract of GG and ZEO (mg GAE g-1 DM).
Total flavonoid contents of GG and ZEO were determined using Aluminium chloride colorimetric assay with quercetin as a standard (Marinova et al., 2005). Briefly, 0.1 ml of each extract (0.1 mg mL-1) or standard quercetin solutions (0–0.5 mg) were separately mixed with 0.3 ml of 5% NaNO2 for 5 min. Then 0.3 ml of 10% AlCl3 was mixed for 6 min and 2 ml of 1 M NaOH was added. The absorbance of the reaction mixtures was measured at 510 nm using a UV-VIS spectrophotometer (2550, Shimadzu, Japan). Total flavonoid contents were expressed as mg of quercetin equivalent per gram of GG and ZEO extracts. Analyses were done in triplicate.
2.7. DPPH photometric assay
Antioxidant activity of GG and ZEO was determined using the 2,2-diphenyl-1-picrylhydrazyl (DPPH) photometric assay. From the both samples at different concentrations, 15 μL were added to 5 mL of a 0.004% of DPPH in methanol. The reaction mixture was mixed in the vortex mixer and kept in the dark at the room temperature for an hour. After 60 min keeping of reaction, the absorbance of the mixture was read against a blank at 517 nm by a UV-VIS spectrophotometer (2550, Shimadzu, Japan). Inhibition percentage of DPPH was calculated according to the following formula:
Inhibition of DDPH (%) =100×(A1 – A0)/A0
Where A0 and A1 are the absorbance of the controls and samples, respectively. Ascorbic acid was used as a positive control.
2.8. Edible coating preparation of Alg/glycerol/GG/ZEO
The central composition of active edible coatings was designed with three factor viz. Alg, GG, and ZEO. The Alg and GG solutions were prepared by dissolving 30 g of sodium alginate and galbanum gum in 1 L of sterile distilled water at 70°C. To prepare the composite film of Alg and GG (CAG), 15 g of Alg and 15 g of GG were blended and developed by the same method. After preparation of solutions, they were stirred at 30 °C for 60 min, then the viscous solutions were left to cool to 20 °C and 0.1 mL of glycerol monostearate (1.0%) (Sigma) was added as a plasticizer to improve the strength and flexibility of the viscous solutions. The pH of the solutions was adjusted to 5.6 with 1 N NaOH. For active edible coating, the different concentrations of ZEO (0, 0.5 and 1%) were mixed with Alg, GG and CAG solutions under magnetic stirring at 55°C. The final solution was homogenized with Ultra-Turrax (Ultra-Turrax, Staufen, Germany) at 7000 rpm for 2 min. The details of the experimental plan with real and coded factors were shown in Table 1.
2.9. Fillet coating
Skinned chicken fillets were prepared from a production line in the same day and packed in polypropylene trays, and then were transferred and kept under refrigeration conditions (4 °C). After preparation of homogeneous 50 mm diameter medallions of 5 mm thick by a stainless steel punch, the fillets were subdivided into two groups: 1) the group was without inoculated pathogenic bacteria, 2) inoculated with L. monocytogenes. In the inoculated group, the fillets were placed in stomacher bags, inoculated with ca. 105 CFU g-1 and homogenized by a stomacher (BagMixer 400, Interscience, France).
Chicken fillets were dipped in the coating solutions based on designed groups for 30 s. Then, the coated fillets stood for 2 min, followed by a second immersion in CaCl2 (Sigma-Aldrich Chemical Co.) for 30 sec to achieve better crosslinking. The coated samples were allowed to drain completely in ambient condition for about 1 h. The samples were placed in sterile polyethylene bags and stored at 4±1°C until testing. The microbiological and chemical evaluation of fillets were conducted at intervals of three days until day 12. The controls were treated similarly in water solution lacking coating materials (Lee & Mooney, 2012).
2.10. Microbiological analysis
Samples were taken for microbial analysis on days 0 (after dipping treatment), 3, 6, 9, and 12 days of refrigerated storage. Microbial counts were performed by methods using plate count agar (PCA) for psychrotrophic and aerobic mesophilic bacteria (AMB), de man-rogosa-sharpe (MRS) agar for lactic acid bacteria (LAB), Pseudomonas agar for Pseudomonas spp., Palcam agar for L. monocytogenes, Violet Red Bile Glucose (VRBG) agar for Enterobacteriaceae family, and dichloran rose bengal chloramphenicol (DRBC) agar for molds and yeasts (MY). Chicken samples (25 g) were aseptically taken in 225 mL of peptone water (0.1%), mixed in a sterile stomacher bags, and homogenized with Stomacher at 200 rpm min-1 for 1 minute. For each sample, appropriate decimal dilutions were serially prepared in tubes containing peptone water (0.1 g 100 ml-1). The inoculated plates were incubated at 37°C for 2 days for total viable counts, Pseudomonas spp., L.monocytogenes, and Enterobacteriaceae. The incubation condition was 7°C for 10 days for psychrotrophic counts, 30°C for 2 days for LAB, and 25°C for 5 days for MY. All the plates were examined visually for the characteristics of a colony (shape, size, pigmentation, etc.) associated with each growth medium. Microbial colonies were counted and expressed as log10 CFU g-1 chicken fillet (Babuskin et al., 2014).
2.11. Determinations of thiobarbituric acid reactive substances (TBARS)
The TBARS was determined colorimetrically as described by Zhang, Wu, and Guo (2016) with some modification. Briefly, 5 g meat was first mixed with 25 mL of trichloroacetic acid 7.5% and 1 mL of 0.5% BHT in ethanol and homogenized. Then, the mixture was macerated with a glass rod and allowed to stand for 1 h at ambient temperature (at 25°C). Next, the mixture was centrifuged at 3600g for 20 min, and the supernatant was filtered through Whatman # 4 filter paper. Afterward, 5 mL of the new filtrate was taken to mix with 5 mL (0.02 M) aqueous solution of 2-thiobarbituric acid (TBA) in a stoppered test tube, then heated in a boiling water bath for 30 minutes, and subsequently cooled to room temperature. The absorbance of cooled samples was measured at a wavelength of 532 nm by a UV-VIS spectrophotometer (2550, Shimadzu, Japan) against a distilled water blank. The TBARS was measured based on malondialdehyde (MDA) mg kg-1 of chicken meat sample.
2.12. Measuring total volatile base bitrogen (TVBN)
To measure TVBN, 10 g of sample was mixed and homogenized with 190 mL of distilled water into a 500-mL round bottom flask. 2 g MgO and one drop of silicone as antifoam solvent were added to the mixture before steam distillation. A 250 mL Erlenmeyer flask containing 25 mL of 3% aqueous solution of boric acid, 0.04 mL of methyl red and methylene blue as mixed indicators for the titration of ammonia was used as the distillate receiver. Due to the resulting TVB-N, the boric acid solution became green. Titration was performed with hydrochloric acid solution (0.1 N) described as TVBN mg per 100 g of chicken fillet. TVBN was calculated as follow:
where, V and C represent the volume of hydrochloric acid and its concentration, respectively (Goulas & Kontominas, 2005).
2.13. Measuring peroxide value
Lipids were extracted according to the method described by Boselli, Velazco, Caboni, and Lercker (2001). In order to measure the peroxide according to AOAC (2005), the extracted oil was dissolved in 30 ml of acetic acid/chloroform (3:2 v/v) and 0.5 ml saturated potassium iodide was added and the mixture was shaken vigorously. After 3 min, the distilled water (30 ml) was added to the mixture and titrated with 0.01N Na2SO4 until a light-yellow color appeared. 0.5 mL Starch solution (1%) was used as a color indicator and titration continued until the blue color disappeared and the light color appeared. This test was done in triplicate for all samples. The peroxide value is then calculated by the formula: (volume of titration sample [ml])×(molarity of thiosulfate [M])× 1000/(weight of sample [g]). The PV value is measured in milliequivalents of oxygen per kilogram of the sample (meq kg-1).
2.14. Sensory evaluation
A panel of 10 trained panelists was selected among the staff of the Nutrition and Food Science Research Center (Science and Research Branch, Islamic Azad University) on the basis of their experience in the sensory analysis. The uncoated/coated fillets after cooking in the microwave at 185C were evaluated based on taste, odor, color, texture, and overall acceptability attributes. The results were expressed on a 9-point hedonic scale. The sensory scores were 9, like extremely; 8, like very much; 7, like moderately; 6, like slightly; 5, neither like nor dislike; 4, dislike slightly; 3, dislike moderately; 2, dislike very much; 1, dislike extremely (Hamedi et al. 2014). The Sensory evaluation of samples was done after 3 days of storage.
2.15. Statistical analysis
Experiments were done twice on different occasions with chicken fillet samples. All analyses were run in triplicate for each replicate. Analysis of all data was performed by One-way analysis of variance (ANOVA) and Duncan’s New Multiple Range Test in SAS version 9.1. The statistical significance of differences between mean values was proved at p<0.05.
3. Results and Discussion
3.1. Chemical analysis of galbanum gum
The general chemical compositions of F. gummosa galbanum are summarized in Table 2. The most abundant monosaccharides were galactose (67.3%), arabinose (18.2%), and uronic acids (14.5%). According to results obtained by Jalali, Ebrahimian, Evtuguin, and Neto (2011) F. gummosa oleo-resin contains the ethanol soluble terpenes and terpenoids (67%), organic solvents insoluble arabinogalactan (25%), and inorganic substances (1.1%). The sugar analysis by same authors revealed 69, 17 and 14% for arabinose, galactose and uronic acids, respectively (Jalali et al., 2011). The difference in results could be explained by the plant growing conditions and different extraction process. Alkaloids and phenol compounds were not found in F. gummosa exudates, but saponin and tannin were found in trace. The predominant fractions of F. gummosa oleo-resin were terpenes and terpenoids were (55%), sesquiterpenes/sesquiterpenoides (30%) and monoterpenes/monoterpenoides (15%). The prominent terpenes and terpenoids of F. gummosa oleo-resin were (+) Norinine, (+) Eremorphilene, Limonene, and β-Amyrin (Jalali et al., 2011).
3.2. Chemical compounds of Ziziphora essential oil (ZEO)
The essential oil from aerial parts of Z. persica was analyzed by GC-MS and resulted in the identification of 47 compounds representing 97.43% of the oil. The yield oils (yellow liquids) was 1.05% (v/v), based on dry weights. The most representative compounds of ZEO were monoterpene and among them, the main constituents were (+)- Pulegone (31.42%), Neomenthol (18.58%), p-Menthone (17.13%), 1,8-Cineole (6.98%) and β-Pinene (2.93%) (Table 3). The result of ZEO analysis was in accordance with other earlier studies on Ziziphora species that all found to be rich in Pulegone (Dembitskii, Bergaliev, & Kyazimov, 1994; Khodaverdi-Samani et al., 2015; Ozturk & Ercisli, 2006; Rustaiyan, Jamzad, Masoudi, & Ameri, 2006). Other Ziziphora species revealed that Pulegone was the main compound of the essential oils of Z. clinopodioides (Shahbazi, 2015), Z. taurica ssp. cleonioides (Meral, Konyalioglu, & Ozturk, 2002), and Z. persica (Rustaiyan et al., 2006). Compared to results of Ozturk and Ercisli (2006) and Nadaf, Halimi, and Nasrabadi (2013), the concentrations of Pulegone, Limonene and Piperitenone of this studies were less than those reported for Z. persica, but these findings are consistent with Rustaiyan et al. (2006). The difference in composition could be influenced by local, climatic, seasonal and experimental conditions.
3.3. MIC and MBC of Ziziphora and galbanum gum
As shown in Table 4, the essential oil of Ziziphora persica proved to be effective against the test microorganisms. Overall, the results of MIC and MBC values ranging from 1.25 to 45.00 (µg mL-1), which all strains were inhibited by ZEO, with different degrees of inhibition. Regarding the MIC and MBC values, B. cereus was the most sensitive strain, while P. aeruginosa was the most resistant to growth inhibition by ZEO, showing the highest determined MIC (37.50 µg mL-1). In general, the tested Gram-positive bacteria were more susceptible to ZEO than Gram-positive bacteria. This phenomenon was reported in other studies and could be attributed to the presence of hydrophobic LPS (lipopolysaccharide) in outer membrane structure of Gram-negative bacteria which is especially impermeable to essential oil molecules and the intrinsic tolerance of bacteria, as well as the nature and combinations of phytochemical compounds present in the essential oils (Lv, Liang, Yuan, & Li, 2011; Silva & Domingues, 2017). The antimicrobial activity other species of Ziziphora (Lamiaceae family) include Z.clinopodioides (Ozturk & Ercisli, 2007), Z. tenuior (Aliakbarlu & Shameli, 2013; Celik, Tutar, Karaman, Hepokur, & Atas, 2016), and Z. capitata (Egamberdieva, Wirth, Behrendt, Ahmad, & Berg, 2017) have been reported.
The MIC and MBC of F. gummosa galbanum gum were determined against selected pathogens (Table 4). In overall, it showed that Gram-positive bacteria have the lowest resistance to Galbanum gum. Data indicated that the lowest inhibitory concentration of GG was 1.25 and 1.50 µg ml-1 belonged to B. cereus and S. aureus, respectively. The antimicrobial activity of GG against Acinetobacter clinical isolates was reported by Afshar et al. (2016). The same result was reported by Abedi, Jalali, and Sadeghi (2009) which indicated the antimicrobial activity of oleogum resin of Ferula gumosa Bioss. essential oil using Alamar Blue™.
3.4. Antioxidant activities of GG and ZEO
The active compounds such as flavonoids, phenolics, curcumanoids, xanthons, tannins, coumarins, lignans, and terpenoids are widely found in food products derived from plant sources and they have been shown to possess significant antioxidant activities (Roby, Sarhan, Selim, & Khalel, 2013). The results for total phenolic, total flavonoid contents and DPPH assay of GG and ZEO extracts are presented in Table 5. Ferula gummosa GG with total phenolic and flavonoid contents (44.13 and 14.26 mg equivalents to gallic acid and quercetin, respectively) showed moderate antioxidant power in DPPH assay. The IC50 (half maximal inhibitory concentrations) for scavenging the 1, 1-diphenyl-2-picrylhydrazyl (DPPH) radicals was 542 μg mL-1. Similar result with a little difference was reported by Ebrahimzadeh, Nabavi, Nabavi, and Dehpour (2011) in the hydroalcoholic extract of Ferula gummosa Boiss roots.
The results of total phenolic, total flavonoid contents and free radical-scavenging activity assay of Z. persica essential oil were summarized in Table 5. Regarding our results from bioactivity evaluation and chemical analysis of ZEO (Table 3), there are different secondary metabolites including terpenoids, flavonoid, and phenolics. These phenolic and flavonoid compounds, especially terpenoides such as menthol, thymol, carvacrol, pulegone, piperitenone, p-menth-3-en-8-ol and 1, 8-cineole are the most important antioxidant components and show strongly radical scavenging effect (Aliakbarlu & Shameli, 2013; Mazandarani & Bovanloo, 2015).
3.5. Antimicrobial activities of edible coatings
The microbial changes of chicken fillet wrapped with Alg, GG, and CAG incorporated with ZEO (0.5 and 1%) films during 12 days of storage at refrigerated temperature are shown in Fig. 1. The number of bacteria, as well as yeast and molds in all samples, increased during 12 days of cold storage time but the value increased faster for control. In general, compared to control samples, all films in the experiment showed significant antimicrobial effects against aerobic mesophilic, psychrotrophic, Pseudomonas spp., LAB, L.monocytogenes, and Enterobacteriaceae, and MY (except Alg coating group that had not significant effect on treated samples in comparison with control groups). This result was in an agreement with Azarakhsh, Osman, Ghazali, Tan, and Adzahan (2014), but some studies indicated that coating with Alg cause a significant decrease in microbial count in some products like as rainbow trout, bream, and bighead carp fillets, (Hamzeh & Rezaei, 2012; Heydari, Bavandi, & Javadian, 2015; Song, Liu, Shen, You, & Luo, 2011). Moreover, the results showed that the fillet treated with GG incorporation with ZEO at the concentration of 1% had insignificantly (P>0.05) lower microbial counts. The initial aerobic mesophilic and psychrotrophic values of control groups were 3.2 and 2.8 and gradually increased to 8.5 and 7.2 log CFU g-1, respectively, at the end of storage. As shown in Fig.1(A and B) the aerobic mesophilic and psychrotrophic values of the treated samples with each of the GG/ZEO (0.5 and 1%), CAG/ZEO (0.5 and 1%), and Alg/ZEO (1%) were below 7 log CFU g-1 until the 12 day that were upper microbiological limit for acceptable quality meat (ICMSF, 2011). With respect to the use of Ziziphora essential oil, present results are in agreement with those of Shavisi, Khanjari, Basti, Misaghi, and Shahbazi (2017), who reported a reduction in bacterial count of minced beef by 2–3 log CFU g-1 (p<0.05) with the addition of 1 and 2 % of Ziziphora clinopodioides essential oil to cellulose nanoparticle film. They are also in agreement with those of Kakaei and Shahbazi (2016) who reported a reduction of total mesophilic and psychrotrophic bacteria in minced beef meat by 2-3 log CFU g-1 when Z. clinopodioides essential oil was added at the concentration of 1 and 2% to chitosan-gelatin film.
The initial LAB counts (Fig. 1.C) were ca. 3.1 log and reached 7.1 log CFU g-1 on day 12 of storage for control. On the same day of storage, the use of GG the films containing ZEO (1%) resulted in a reduction in LAB counts by almost 2.9 log CFU g-1 (p<0.05), while the concentration of 1% of ZEO with Alg and CAG edible coatings decreased the LAB count in fillet by 1.5 and 2.5 CFU g-1, respectively. The results also demonstrated that GG film was more effective than CAG and Alg coating in reducing the populations of LAB.
Enterobacteriaceae and coliforms are usually considered by food manufacturers as hygiene indicators of sanitary quality of foods and water and substantial group of the chicken meat microbial flora. In our study, the initial log of Enterobacteriaceae was recorded to be 2.1, and it was reached to 7.5, 7.2, 5.8 and 6 logs on day 12 in control, Alg, GG and CAG groups, respectively (Fig. 1.D). The incorporation of ZEO into different edible coating caused a significant reduction of Enterobacteriaceae count as compared to control samples (p>0.05). Similar results were observed by Raeisi, Tajik, Aliakbarlu, Mirhosseini, and Hosseini (2015), who found a reduction of 1–2 log cycles in Enterobacteriaceae count by using alginate coating with nisin, cinnamon, and rosemary EOs on microbial quality of chicken meat. Also, the in vitro antimicrobial activity of the Ziziphora and Ferula gummosa EOs against Enterobacteriaceae was reported by Shahbazi (2016), and Ghasemi, Faridi, Mehregan, and Mohagheghzadeh (2005).
The changes of Pseudomonas spp. count in treated groups was similar to aerobic mesophilic and psychrotrophic values (Fig. 1.E). The initial count of Pseudomonas spp. was 2.3 log CFU g-1 and the final population in the untreated sample was up to the level of 7.3 log CFU g-1 (day 12). Raeisi et al. (2015) reported that alginate coating with cinnamon and rosemary EOs had a significant effect on Pseudomonas spp. in chicken fillet during cold storage time, which is in good agreement with our findings. In another study, Gómez-Estaca, De Lacey, López-Caballero, Gómez-Guillén, and Montero (2010) found that biodegradable gelatin–chitosan films containing EOs of clove, fennel, cypress, lavender, thyme, herb-of-the-cross, pine, and rosemary exerted an inhibitory effect on the Pseudomonads count of fish patties.
The initial count of mold and yeast of fillet was very low (less than 1.4 log CFU g-1), however, during 12-day storage, it increased to 5.6 and 5.3 log CFU g-1 for control and Alg samples, respectively. As expected, in general, adding the ZEO to coated samples cause lower counts than the controls (p<0.05), and the lowest values of MY count belonged to GG containing 1% ZEO with 2.1 log CFU g-1 on day 12. Results of this study showed galbanum gum had a notable antifungal activity that caused growth inhibition of MY in chicken fillet samples (Fig. 1.E). Therefore, fungi and yeast showed more sensitivity to F. gummosa GG than the bacteria. These results also were in agreement with Raeisi et al. (2015) who found that the sodium alginate with rosemary and cinnamon EOs caused a significant reduction of MY counts. Overall, the active edible films containing EOs are, in general, very effective against yeasts and molds. This may be due to that the edible coating act as a barrier for oxygen transfer leads to inhibition of molds (Heydari et al., 2015).
The effect of Alg, GG, CAG and their combination with ZEO for the control of L. monocytogenes as a function of storage time is given in Fig. 2. According to results, no significant differences (p>0.05) was observed between the growth of L. monocytogenes in control and Alg coated samples during storage (Fig. 2). These findings indicate that the Alg coating had no significant inhibitory effect on the growth of L. monocytogenes (similar to Pseudomonads, LAB, and Enterobacteriaceae ), but samples treated with GG, CAG, separately and in combination with ZEO, reduced the growth of L. monocytogenes ( Fig. 2). ZEO contains high concentration of monoterpenes and phenolic compounds, and their synergistic effect can enhance the antimicrobial activity of active edible coating against the foodborne pathogens (Kakaei & Shahbazi, 2016). After 12 days of cold storage, the inoculated L. monocytogenes count reached a value of 6.8 log CFU g-1 in the control samples, while the use of edible Alg, GG, and CAG enriched with the highest concentration of Ziziphora maintained the population of L. monocytogenes under the 4.3, 2.1 and 2.7 log CFU g-1, respectively. In a study conducted by Alboofetileh, Rezaei, Hosseini, and Abdollahi (2014) the antibacterial effects of clove, coriander, caraway, marjoram, cinnamon, and cumin essential oils were incorporated into alginate-clay nanocomposite film were evaluated against L. monocytogenes, E. coli, and S. aureus. Their results showed that the combination of EO with alginate was effective to control the growth of pathogens in culture media (in vitro). The same result was reported by Benavides, Villalobos-Carvajal, and Reyes (2012) in oregano–alginate-based film against the aforementioned strains.
3.6. Chemical analysis of coated fillet
The measurement of TBARS is a well-established method for screening and monitoring lipid peroxidation and has been widely used to estimate the extent of lipid oxidation and sensory evaluation in meat product like as chicken fillet (Xiong et al., 2015). Also, peroxide value and total volatile basic nitrogen (TVBN) are widely used measurements of meat and meat products quality. The results of TBARS, peroxide and TVBN values for treated samples are presented in Fig. 3(A). The TBARS formation in the samples treated by GG and CAG alone and in combination with ZEO, as well as Alg/ZEO were significantly lower than the control group (p<0.05). There was no significant difference recorded between Alg and control samples during 12 days of cold storage (0.604 and 0.586 mg MDA kg-1, respectively). Samples coated by GG/ZEO had less increase of TBARS in respect to other treatments during storage at 4˚C (0.259 mg MDA kg-1). Vital et al. (2016) evaluated the effects of an alginate-based edible coating containing rosemary and oregano EOs on lipid oxidation, color preservation, water losses, texture and pH of beef steaks during 14 days storage. The reported TBARS values for this study were reached approximately 1.00, 0.91, 0.61 and 0.53 mg MDA/kg for control, Alg, Alg/rosemary and Alg/oregano samples, respectively. The in vitro antioxidant properties of some EOs especially Ziziphora and Ferulagummosa were reported by Ghafari et al. (2006) and Ebrahimzadeh et al. (2011). In another study, Song et al. (2011) showed that alginate-based edible coating with vitamin C and tea polyphenols had lower TBARS values in comparison with the uncoated samples. So active edible coating incorporating a natural antioxidant, such as ZEO, may prolong the shelf life of meat products by its antioxidative activity.
The initial TVBN values of uncoated fillet samples were 13.7 and reached 42.95 mg N 100 g-1 on day 12 (Fig. 3.B). TVBN increase in control samples was expected because it is related to bacterial spoilage (Silva and Domingues, 2017), but the high amount of TVBN in Alg coated group can be attributed to its low antimicrobial activity. In contrast, Lee and Mooney (2012) reported that the TVBN content of bighead carp fillets coated with sodium alginate was significantly lower than the control group in the last 8 days of the storage period (p<0.05). The TVBN value of GG and CAG with 1% ZEO samples remained practically below acceptable values (20.63 and 25.65 mg N 100 g-1, respectively) and this is associated with the higher antimicrobial effect of GG incorporation with ZEO.
As seen in Fig. 3(C), the PV of the fillets increased gradually in all treatments during the period, but this value in all samples was not significantly different in first 3 days of our investigation. The initial PVs were very low (average 0.0135 meq kg-1) in the fresh fillets, and this value in specimens containing GG, CAG, GG/ZEO and CAG/ZEO was less than control and Alg groups on 9 and 12 days of storage (Fig. 3.C). The present study showed that active edible coating with GG and ZEO delayed oxidation of fillet better than other samples.
All samples were evaluated based on 9-point hedonic scale, and the score of 7 or more considered satisfactory. The summary of the sensory evaluation includes color, odor, taste, texture and overall acceptance are given in Fig. 4. As the results show the control, Alg, and Alg/ZEO 0.5% coated samples had a high value of taste and overall acceptance parameters and there was no significant difference between them. The higher score of color and texture belonged to GG and GG/ZEO samples, and the value of samples coated by Alg and CAG-containing 1% ZEO were still in acceptable ranges. Kulig, Zimoch-Korzycka, Król, Oziembłowski, and Jarmoluk (2017) indicated that sensory properties of raw and cooked meat samples coated with Alginate/Chitosan polyelectrolyte complex had no undesirable influence on pork meat products. (Vital et al. (2016)) also suggested that an alginate-based edible coating containing natural antioxidants (rosemary and oregano EOs) had a significant effect on consumer perception of odor, flavor and overall acceptance of the beef.
The present study showed that sodium alginate, galbanum gum and composite coating (CAG) enriched the chemical and microbial properties of chicken fillet and increased its shelf life. The combination of these coatings with ZEO showed significantly higher in vitro antioxidant and antimicrobial activities and they had a high chemical and microbial quality than uncoated fillets. Antioxidant and antibacterial effects of Alg, GG, and CAG coating with ZEO were more pronounced when the essential oil was used at high concentration. Therefore, alginate and new edible films that used in this study (GG and CAG), alone and in combination with other natural active products like as essential oils, can be used for to improve food quality and enhance the shelf life of perishable foods like as meat and meat products.
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Fig. 1: Antimicrobial effects of active edible coating by sodium alginate (Alg), galbanum oleo-resin gum (GG), composite film of alginate and galbanum (CAG) incorporation with Ziziphora essential oil (ZEO) in chicken fillet during 12 days storage at 4°C. (A): Total aerobic mesophilic bacteria, (B): Total psychotrophic bacteria, (C): Lactic acid bacteria, (D): Enterobacteriaceae family, (E): Pseudomonas spp., and (F): molds and yeasts counts.
Fig. 2: Listeria. monocytogenes bacteria count in chicken fillet coated by sodium alginate (Alg), galbanum oleo-resin gum (GG), composite film of alginate and galbanum (CAG) incorporation with Ziziphora essential oil (ZEO) during 12 days storage at 4°C.
Fig. 3: The chemical changes of chicken fillet coated by sodium alginate (Alg), galbanum oleo-resin gum (GG), composite film of alginate and galbanum (CAG) incorporation with Ziziphora essential oil (ZEO) during 12 days storage at 4°C. (A): TBARS, (B): TVBN, and (C): Peroxide values.
Fig. 4: Sensory evaluation of samples coated sodium alginate (Alg), galbanum oleo-resin gum (GG), composite film of alginate and galbanum (CAG) incorporation with Ziziphora essential oil (ZEO) during 12 days storage at 4°C.
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