Olive leaf extract and usage for development of antimicrobial food packaging
Z.Ö. Erdohan and K.N. Turhan,
Department of Food Engineering, University of Mersin, 33343, Çiftlikköy, Mersin, Turkey.
In recent years, studies on usage of antimicrobial plant extracts, like thyme, sage and olive leaf extracts, have gained acceleration since these are generally classified as GRAS (generally recognized as safe). As an important part of these studies is focused on medical beneficences, the rest of them are focused on alternative usage of the extracts. Antimicrobial packaging applications are really interesting field among these studies. This part of active packaging prolongs shelf life and maintains quality and safety of food products by extending the lag phase and reducing the growth rate of food spoilage microorganisms or food pathogens. This chapter reviews antimicrobial efficiency of olive leaf extract against some food pathogens and usage of this extract in antimicrobial packaging materials and inhibitory efficiencies against Staphylococcus aureus.
The knowledge of the medicinal properties of the olive tree (Olea europaea) date back to the early 1800's where it was used in liquid form as a very effective treatment for malarial infections. Pulverized leaves were used in a drink to lower fevers and a few decades later, green olive leaves were used in tea as a treatment for malaria [1]. According to Pharmaceutical Journal of Olive (1854), olive leaves have been recognized for providing many benefits in food as well as cosmetic and mostly nutraceutical applications. Now, olive leaf extract (OLE) is known with its high antioxidant, antimicrobial and antibacterial activity. OLE is very effective activity against various diseases, such as coronary artery disease, hypertension, high cholesterol level, arrhythmia, cancer, diabetes, overweight, osteoporosis, herpes, flu and colds, and some bacterial, fungus and yeast infections. It is also a natural, genetically modified organism free and allergen free product. Thus, OLE has been served in capsule form to make it easily taken recently [2]. The antioxidant and antimicrobial efficiency of the OLE are directly related with its polyphenols. There are some studies in literature revealing that polyphenols can inhibit the sporulation of Bacillus cereus and growth of Escherichia coli, Klebsiella pneumoniae, Salmonella typhi, Vibrio parahaemolyticus and Staphylococcus aureus, that all known as food pathogenes. To prevent of these pathogens, heat treatments, cold pasteurization techniques, addition of preservatives and some antimicrobials in food formulation or spread these additives on food surfaces may be applied. Antimicrobial packaging is an innovative way of inhibiting microbial growth on the foods while maintaining quality, freshness, and safety. Although there has been a rising interest in the researches in this field, availability of antimicrobials and new polymeric materials, regulatory concerns, and appropriate testing methods limit the developments. Antimicrobial packaging is highly regulated around the world and researchers must take these regulations into consideration [3].
2. Effects of Olive Leaf Extract on Food Pathogens
Olive and its products are an important part of the Mediterranean diet and olive leaf is the by-product of the olive oil industry. The leaves are rich in polyphenols, namely oleuropein, tyrosol, hydroxytyrosol, rutin, verbacoside, apigenin-7-glucoside and luteolin-7-glucoside [4, 5]. There are some studies in the literature showing that polyphenols are responsible for the functional properties especially antimicrobial activity [5-8]. In these studies, the phenolics are extracted with different solvents, and isolation or purification can also be applied for specific one. The choice of solvents used in extracts, cultivars of olives, crop origin, harvesting time and climate may all change the leaf composition, which could influence antibacterial activities of extracts [5, 9]. The solvent type is the most important factor affecting the efficiency of liquid-solid extraction [10]. There are many studies in the literature showing the effect of solvents used in extraction on the phenolics distribution and total phenol content in the OLEs [6, 10, 11]. An HPLC chromatogram of phenolics in the chloroform/methanol extracted OLE is shown in Figure 1 and the phenolic compounds distributions in different OLEs are given in Table 1.
Science against microbial pathogens: communicating current research and technological advances A. Méndez-Vilas (Ed.)______________________________________________________________________________
min0 5 10 15 20 25 30 35 40
mAU
0
500
1000
1500
2000
DAD1 A, Sig=280,4 Ref=400,100 (SIG00610.D)
1
23 4
5 6
7
8
10
12
Figure 1 HPLC chromatogram of phenolics in the chloroform/methanol (50/50, v/v) extracted OLE [9]. Oleuropein is the most abundant compound in OLEs given in Table 1 and followed by apigenin-7-glucoside for Extract A; hydroxytyrosol for Extract B and Extract D, and verbascoside for Extract C. Difference in the phenolics distribution may caused by the solvent type with the other factors mentioned above. Altıok and others (2008) examined the effect of different solvents on the total phenolic content of OLEs and oleuropein abundance in extracts. They obtained an extract with highest total phenolics (10.3 mg total phenolics/g leaf) and oleuropein (92 mg oleuropein/g leaf) content using 70% ethanol [10]. Pereira and others (2007) used water as solvent to obtain OLE and they detected 26.5 mg oleuropein/g lyophilized olive leaves extract [6]. Le Floch and others (1998) compared phenol yields of OLE obtained by supercritical fluid extraction and sonication. They used methanol, n-hexane, diethyl ether, ethyl acetate and carbon dioxide modified with 10% methanol as solvent for extraction. They obtained the highest and lowest phenol yields with methanol and n-hexane extraction, respectively. They also revealed supercritical fluid extraction was much successive than sonication extractions with n-hexane, diethyl ether and ethyl acetate [11].
Table 1 Peak numbers and abundances (peak area, %) of the main phenolic compounds present in OLE solutions [1, 4, 10].
Peak no Phenolic Compounds Peak Area (%)
Extract A [4] Extract B [4] Extract C [10] Extract D [1]
Science against microbial pathogens: communicating current research and technological advances A. Méndez-Vilas (Ed.)_______________________________________________________________________________
Table 2 Antimicrobial efficiency of OLEs extracted with different solvents against S. aureus [9].
Solvent Inhibition Zone Diameter (mm) Water 29.4±1.1 a Ethanol 24.8±0.7 c Methanol 25.3±1.0 c Chloroform 16.7±0.7 d Chloroform/Ethanol (50/50, v/v) 27.3±1.2 b Chloroform/Methanol (50/50, v/v) 27.1±0.9 b
Different superscript letters in each column are significantly different (p<0.05). Difference in the phenolic compounds distribution strongly affects the functionality of the extracts such as antimicrobial activity. Turhan (2009) investigated the effect of different solvents used in extraction on the antimicrobial activity of OLEs against Staphylococcus aureus by paper disc bioassay (Table 2). It was reported that the concentration of 129 mg/mL olive leaf water extract showed the highest inhibitory effect on S. aureus and followed by chloroform/ethanol and chloroform/methanol extracts [9]. Antimicrobial efficiency of phenolics extracted from olive, olive oil and olive leaf against some food pathogens, namely S. aureus, E. coli, S. enteritidis and S. thypimurium, Listeria monositogenes are given in Table 3. Markin and others (2003) investigated the minimal bactericidal concentration of olive leaf water extract against S. aureus, B. subtilis, P. aeruginosa, K. pneumoniae and E. coli. They defined the minimal bactericidal concentration of the OLE as 6 mg/mL for S. aureus and E. coli (Table 3). They observed a complete destruction of S. aureus within 2 h. Moreover increasing OLE concentration from 3 mg/mL to 6 mg/mL caused a decrease the inhibition time from 24 h to 3 h for E. coli [14]. In another study, Pereira and others (2007) tested the antimicrobial activity of aqueous olive leaf extract against S. aureus, B. subtilis, P. aeruginosa, E. coli, K. pneumoniae and Bacillus cereus, bacteria, Candida albicans and Candida neoformans, fungi. They revealed that the growth rates of S. aureus and E. coli were decreased while OLE concentration increased and the OLE showed a IC25 (25% inhibitory concentration) value of 2.68 and 1.81 mg/mL for S. aureus and E. coli, respectively. Moreover increasing OLE concentration from 0.05 mg/mL to 5 mg/mL caused a significant decrease in optical densities of OLE added S. aureus and E. coli cultivars decreased 51% and 46%, respectively [6]. In another study, Korukluoglu and others (2010) investigated the effect of the extraction solvent on the antimicrobial efficiency of S. aureus, E. coli, S. enteritidis, S. thypimurium and some others. They reported that solvent type affected the phenolic distribution and concentration in extracts, and antimicrobial activity against tested bacteria. As ethanol extracted OLE showed the highest antimicrobial efficiency against E. coli and S. enteritidis, acetone extracted OLE showed the highest antimicrobial efficiency against S. thypimurium [5]. In some studies, the individual antimicrobial effects of olive leaf phenolics were compared with natural OLEs which are mixtures of these phenolics. Results showed that the extracts may be more beneficial than isolated constituents since a bioactive component can change its properties in the presence of other compounds. In another words, a synergistic antimicrobial effect is observed when the phenolic compounds are used together [6]. Considering all effects of OLE against the food borne pathogens, are mentioned above, OLE has a great potential to use as antimicrobial food additive or antimicrobial agent for food packaging.
3. Antimicrobial Packaging
Microbial growth on the product surface is the main cause of spoilage of many foods. To control undesirable microorganisms in foods during storage and distribution, antimicrobial substances can be either coated onto the food surface or incorporated into the food packaging materials [17]. As antimicrobial packaging films are used for surface preservation, antimicrobial containers and utensils are used for liquid foods such as pasteurized egg white or fruit juice. The application of antimicrobial agents to packaging can create an environment inside the package that may delay or even prevent the growth of microorganisms on the product surface by extending the lag period and reducing the growth rate of the microorganisms [17, 18]. Hence, it leads to an extension of the shelf life and/or the improved safety of the product. There are many studies showing the efficiency of the antimicrobial packaging on food pathogens like S. aureus, L. monositogenes, E. coli, S. enteritidis, and S. thypimurium. S. aureus is an important pathogen, due to a combination of toxin-mediated virulence, invasiveness, and antibiotic resistance [19]. Among the food poisoning events, staphylococcal poisoning takes a grade place. In all cases of staphylococcal food poisoning, one of the foodstuffs or ingredients is contaminated with an enterotoxin producing S. aureus strain. Many foods can serve a good growth medium for S. aureus, and have been implicated in staphylococcal food poisoning [19]. Two ways can be followed to prevent staphylococcal poisoning after applying a sufficient thermal process. One way is avoiding from contamination of the food with S. aureus and the second way is inhibition of the microbial growth before the living organism number reaching the critical level if the contamination occurs.
Science against microbial pathogens: communicating current research and technological advances A. Méndez-Vilas (Ed.)______________________________________________________________________________
The efficiency of some antimicrobial packaging materials containing certain antimicrobial agents like nisin, essential oils of garlic, oregano, grapefruit seed and others have been widely investigated (Table 4). These studies underline that a proper polymer with an antimicrobial may offer effective food preservation. However, application of plant extracts or essential oils on a foodstuff or a packaging material may not always reflect the results obtained from preliminary in vitro studies of the same compounds. Foods are complex systems consisting of different connected microenvironments and the antimicrobial additive should release enough from the polymer matrix to show inhibitory effect [18]. For example, rosemary oil showed activity against S. aureus in agar diffusion test but when used in packaging material, they could not reduce the S. aureus counts (Table 4). Thus, having an inhibition effect against certain group of bacteria of an antimicrobial agent is not sufficient alone; selection of an appropriate polymer should be taken account when developing antimicrobial packaging materials.
Table 3 Antimicrobial efficiency of phenolics extracted from olive, olive oil and olive leaf against S. aureus, E. coli, L. monocytogenes, S. enteritidis and S. thypimurium.
Source Solvent Used in Extraction
Inhibition Concentration
Inhibition (%)
Inhibition ZoneDiameter (mm)
Staphylococcus aureus
Olive [12] a Oleuropein hydrolysate [12] a Frozen olives [12] a
Chloroform Ethyl acetate Ethyl acetate
3.5 mg/disc 7.5 mg/disc 10.0 mg/disc
- - -
16-52 47 26
Isolated oleuropein [13] b - 62.5 μg/mL 100 - Isolated hydroxytyrosol [13] b - 7.85 μg/mL 100 - Olive leaf [14] b Water 6 mg/mL 100 - Olive oil [15] c - - 60 - Olive [16] b Methanol 2.84 mg/mL 50 - Olive leaf [6] b Water 2.68 mg/mL 25 -
Olive leaf [5] b Acetone Diethyl ether Ethanol
55 μg/mL 50 μg/mL 55 μg/mL
100 100 100
- - -
Commercial OLE [7] b - 0.78% (v/v) 100 - Escherichia coli
Olive [12] a Chloroform Ethyl acetate Ethyl acetate
3.5 mg/disc 7.5 mg/disc 10.0 mg/disc
- - -
15 - -
Olive leaf [14] b Water 6 mg/mL 3 mg/mL
100 100
- -
Olive [16] b Methanol 0.72 mg/mL 50 - Olive leaf [6] b Water 1.81 mg/mL 25 -
Olive leaf [5] b Acetone Diethyl ether Ethanol
60 μg/mL 72 μg/mL 65 μg/mL
100 100 100
- - -
Commercial OLE [7] b - 25-50% (v/v) 100 - Salmonella enteritidis
Olive leaf [5] b Acetone Diethyl ether Ethanol
170 μg/mL 178 μg/mL 185 μg/mL
100 100 100
- - -
Salmonella typhimurium
Olive [12] b Chloroform Ethyl acetate Ethyl acetate
3.5 mg/disc 7.5 mg/disc 10.0 mg/disc
- - -
16 15 -
Isolated oleuropein [13] b - 125 μg/mL 100 - Isolated hydroxytyrosol [13] b - 3.94 μg/mL 100 -
Olive leaf [5] b Acetone Diethyl ether Ethanol
110 μg/mL 104 μg/mL 90 μg/mL
100 100 100
- - -
Listeria monocytogenes Commercial OLE [7] b - 25% (v/v) 100 -
Science against microbial pathogens: communicating current research and technological advances A. Méndez-Vilas (Ed.)_______________________________________________________________________________
Antimicrobial packaging may be an effective way of food preservation, but there are some critical concerns. First of all, antimicrobial agents may be incorporated into the packaging materials initially or coated or immobilized on the surface of packaging material and migrate into through the food diffusion and partitioning for it to be effective [18]. Thus, this antimicrobial agent must be considered as a food additive and it should correspond to the regulatory concerns [18, 28, 29]. Moreover, as much as antimicrobial efficiency, the cost-to-benefit ratio is an important concern. Some antimicrobial systems can be effective, but if produced on a large scale, they might require expenses beyond the benefits obtained by an extended shelf life or improvement in quality [28]. Lastly, there are numerous technical challenges related to coating methods, the rate of curing, the ease of heat sealing, the effects on physical and mechanical properties of film, the effects on color, the texture or flavor of the food, and the ability of the antimicrobial agent to provide effectiveness throughout the package/product life cycle. The challenges can be daunting, but as research in the field progresses, there is promise for many systems to meet these challenges [3, 28].
Table 4 Inhibition efficiency of some antimicrobial packaging materials against S. aureus.
Antimicrobial Agent
Polymer Application Concentration of Antimicrobial
Inhibition Incubation or Storage Time
Inhibition Zone Diameter/Area
Nisaplin ® [20] CP Cheddar cheese Ham
2560 AU/cm2 1 log CFU/g 2 log CFU/g
12 week 12 week
Garlic oil [21] A NA 0.2 (mL/100 mL) 0.3 (mL/100 mL) 0.4 (mL/100 mL)
4. Antimicrobial Activity of OLE Incorporated Films
Although OLE is known with its antimicrobial activity against several bacteria and fungi, incorporation of OLE in a packaging material as an antimicrobial agent is a new subject. The first study on the OLE incorporated antimicrobial packaging materials and application of it performed by Ayana and Turhan (2009). In the first part of the study, they prepared OLE incorporated antimicrobial MC films and investigated OLE concentration on the film properties (water vapor permeability and mechanical properties) and the antimicrobial efficiency of these films against S. aureus by agar diffusion method. Water extracted OLE was used in a ratio of 0.5-3.0 g OLE/100 mL film solution. OLE concentration in the MC film discs changed within a range of 0.6-3.6 mg OLE/film disc, and increasing concentration of OLE caused a significant increase in the inhibition zone from 16.57 mm to 26.60 mm for S. aureus (Table 5).
Science against microbial pathogens: communicating current research and technological advances A. Méndez-Vilas (Ed.)______________________________________________________________________________
Table 5 The inhibition zone diameters of MC films containing OLE against S. aureus [30].
OLE (g/100 mL)
OLE (mg/film disc)
MC films (mm)
Contact Surface
0 0 0 g + 0.50 0.6 16.57±1.48 f - 0.75 0.9 18.77±1.26 e - 1.00 1.2 21.37±1.41 d - 1.50 1.8 23.97±0.84 c - 2.00 2.4 25.17±1.03 b - 3.00 3.6 26.60±0.21 a -
Different superscript letters in each column are significantly different (p<0.05). Although an antimicrobial material show antimicrobial effect on target microorganisms, effectiveness of this package when applied on a food may be different. Foods have different chemical and biological characteristics and they provide different environmental conditions to microorganisms and included antimicrobial agents. Moreover, packaging materials modify the atmosphere in the package and water activity of the food may change because of water vapor permeability of the packaging materials [18]. Thus food characteristics and packaging material properties should be taking account when an antimicrobial packaging material is applied [30]. In the second part of the study, Ayana and Turhan (2009) applied the OLE incorporated MC films on the S. aureus inoculated kasar cheese and investigated the inhibition efficiency of the films [30]. The water vapor permeability of the packaging material affects water activity which causes microbial spoilage, decreases quality and storage stability of food products. For the application, not only antimicrobial activity but also other properties of the films like water vapor permeability, mechanical properties and sensorial attributes were taken account for the film efficiency. The lowest water vapor permeability and the highest mechanical properties were obtained from the MC films containing 1.5 and 2.0 g OLE/100 mL film solution, respectively. Although increasing concentration of OLE caused an increase on the antimicrobial activity of the film, the transparency of the film decreased and the bitterness increased. Thus, the film containing 1.5 g OLE/100 mL solution was selected to wrap the kasar cheese slices inoculated with S. aureus [30]. They reported that the S. aureus numbers decreased 1.22 log cycle for the slices wrapped with OLE incorporated MC films after 14 day of storage. On the other hand S. aureus count increased 0.64 and 0.60 log cycle at the end of the storage for non-wrapped and MC film wrapped slices, respectively (Table 6). A similar study was performed by Scannel and others (2000) who packed S. aureus inoculated cheddar cheese slices with Nisaplin® immobilized cellulosic pouches. After 14 and 84 day of storage, S. aureus count decreased approximately 0.4 and 1 log cycle, respectively [20].
Table 6 The numbers of S. aureus on kasar cheese slices during storage [30].
Day
S. aureus number (log CFU/cm2)
Non-wrapped MC film wrapped OLE incorporated MC
film wrapped
0 5.31±0.22 c 5.05±0.04 de 4.97±0.28 e 4 5.21±0.08 cde 5.02±0.18 de 4.49±0.12 f 7 5.34±0.03 c 5.27±0.07 cd 4.35±0.10 f
11 5.74±0.05 ab 5.57±0.11 b 3.56±0.14 g 14 5.95±0.13 a 5.65±0.03 b 3.75±0.30 g
Means in same column and row with different superscript (a-g) are significantly different (p<0.05). In another study, antimicrobial efficiency of OLE incorporated PLA and MC-PLA films were investigated against S. aureus [4]. Although water extracted OLE show the highest inhibitory effect against S. aureus [9], the extract was incompatible with PLA and MC-PLA film solutions. Thus, the chloroform-methanol extracted OLE was used for preparing antimicrobial PLA and MC-PLA films. It was reported that the MC-PLA films were much successive inhibiting S. aureus than the PLA films at all OLE concentrations and increasing amount of the OLE in the PLA and MC-PLA film discs caused a significant increase in the inhibitory zones from 9.10 mm to 16.20 mm and from 9.75 mm to 20.15 mm, respectively (Table 7). It was suggested that the incorporation of hydrophilic MC into the hydrophobic PLA matrix probably made easy the release of the OLE to the hydrophilic agar medium. When the S. aureus inhibition efficiencies of PLA and MC-PLA films are compared with MC films, it is seen that MC film much effective than others (Table 5-7). To show inhibitory effect, a non-volatile antimicrobial substance
Science against microbial pathogens: communicating current research and technological advances A. Méndez-Vilas (Ed.)_______________________________________________________________________________
should release from the polymer surface by diffusion. Diffusion of the antimicrobial substance occurs with a mechanism of diffusion of penetrant into the polymer matrix, swelling and dissolution the polymer matrix and release of the antimicrobial molecules from the matrix [31]. There are some studies in the literature showing the effect of hydrophilicity of the polymer matrix on the antimicrobial agents release from the polymer films [32, 33]. According to these studies, as hydrophilic substance incorporation causes an increase on the release of antimicrobials, hydrophobic substance incorporation shows reverse effect. Incorporation of hydrophobic substances into the polymer may restructure the matrix by increasing the network tortuosity, which may affect other geometric features, such as pore constrictions or blind porosity, thereby limiting molecular transport through the network [32]. Moreover, hydrophobic compounds reduce diffusion of hydrophilic compounds release from the film by slowing down film hydration. Thus, inhibitory effect of antimicrobial PLA and MC-PLA films may be lower than the OLE incorporated antimicrobial MC films.
Table 7 The inhibition zone diameters of PLA and MC-PLA films containing OLE against S. aureus [4].
Different superscript letters in each column are significantly different (p<0.05).
Figure 2 The effect of OLE amount on antimicrobial efficiency of PLA (A) and MC-PLA (B) films against S. aureus (ATCC 25923) A1-B1: 0 g OLE/100 mL, A2-B2: 0.5 g OLE/100 mL, A3-B3: 3 g OLE/100 mL [4].
5. Conclusion
Usage of plant extract, like olive leaf extract, as antimicrobial substance is still new subject for antimicrobial food packaging. OLE incorporated MC, PLA and MC-PLA films have a great potential in antimicrobial food packaging to reduce post-process growth of S. aureus and decrease staphylococcal food poisoning events. Moreover, the use of hydrophilic polymers (MC, starch, pectin etc.) with a hydrophobic polymer (PLA, petroleum based polymers etc.) may offer a good way for carrying hydrophilic antimicrobials. Because of the biodegradability, having good barrier and mechanical properties PLA based antimicrobial packaging materials have a potential to commercialize.
OLE (g/100 mL)
OLE (mg/film disc)
Inhibitory Zones (mm) Contact Surface
PLA films MC-PLA films 0 0 0 f 0 f +
0.50 0.9 9.10±0.31 e 9.75±0.85 e - 1.00 1.8 10.75±0.72 d 14.10±1.41 d - 1.50 2.7 13.45±0.51 c 16.15±0.93 c - 2.00 3.6 14.50±0.69 b 18.00±1.62 b - 3.00 5.4 16.20±0.83 a 20.15±0.81 a -
Science against microbial pathogens: communicating current research and technological advances A. Méndez-Vilas (Ed.)______________________________________________________________________________
References
[1] Benavente-Garcõa O, Castillo J, Lorente J, Ortuno A, Del Rio JA. Antioxidant activity of phenolics extracted from Olea europaea L. leaves. Food Chemistry. 2000; 68: 457-462.
[2] Ritchason J. Olive leaf extract-Potent antimicrobial, antiviral and antifungal agent, Woodland Publishing; Australia, 2000. [3] Appendini P, Hotchkiss JH. Review of antimicrobial food packaging. Innovative Food Science and Emerging Technologies.
2002; 3(2): 113-126. [4] Erdohan ZO, Cam B, Turhan KN, Journal of Food Science. 2011 (submitted). [5] Korukluoglu M, Sahan Y, Yigit A, Ozer ET, Gucer S. Antibacterial activity and chemical constitutions of Olea europaea L. leaf
extracts. Journal of Food Processing and Preservation. 2010; 34: 383-396. [6] Pereira AP, Ferreira ICFR, Marcelino F, Valentão P, Andrade PB, Seabra R, Estevinho L, Bento A, Pereira JA. Phenolic
compounds and antimicrobial activity of olive (Olea europaea L. Cv. Cobrançosa) leaves. Molecules. 2007; 12: 1153-1162. [7] Sudjana AN, D’Orazio C, Ryan V, Rasool N, Ng J, Islam N, Riley TV, Hammer KA. Antimicrobial activity of commercial Olea
europaea (olive) leaf extract. International Journal of Antimicrobial Agents. 2008; 33: 461-463. [8] Aytul KK. Antimicrobial and antioxidant activities of olive leaf extract and its food applications. Turkey, Ms. Thesis, Graduate
School of Engineering and Sciences of Izmir Institute of Technology; 2010. [9] Turhan KN. Production and characterization of antimicrobial polylactic acid and methylcellulose-polylactic acid films. The
Scientific and Technological Research Council of Turkey, Turkey. Project No: 107O234, 2009. [10] Altıok E, Baycin D, Bayraktar O, Ulku S. Isolation of polyphenols from the extracts of olive leaves (Olea europaea L.) by
adsorption on silk fibroin. Separation and Purification Technology. 2008; 62(2): 342-348. [11] Le Floch F, Tena MT, Ríos A, Valcárcel M. Supercritical fluid extraction of phenolic compounds from olive leaves. Talanta.
1998; 46: 1123-1130. [12] Fleming HP, Walter Jr. WM, Etchells JL. Antimicrobial properties of oleuropein and products of its hydrolysis from green
olives. Applied and Environmental Microbiolgy November. 1973; 26(5): 777-782. [13] Bisignano G, Tomaino A, Lo Cascio R, Crisafi G, Uccella N, Saija A. On the in-vitro antimicrobial activity of oleuropein and
hydroxytyrosol. Journal of Pharmacy and Pharmacology. 1999; 51: 971-974. [14] Markin D, Duek L, Berdicevsky I. In vitro antimicrobial activity of olive leaves. Mycoses. 2003; 46: 132–136. [15] Medina E, Castro A, Romero C, Brenes M. Comparison of the concentrations of phenolic compounds in olive oils and other
plant oils: Correlation with antimicrobial activity. Journal of Agricultural Food Chemistry. 2006; 54(14): 4954-4961. [16] Sousa A, Ferreira ICFR, Calhelha R, Andrade PB, Valentão P, Seabra R, Estevinho L, Bento A, Pereira JA. Phenolics and
antimicrobial activity of traditional stoned table olives ‘alcaparra’. Bioorganic and Medicinal Chemistry. 2006; 14: 8533-8538. [17] Cha DS, Chinnan MS. Biopolymer-based antimicrobial packaging: A review. Critical Reviews in Food Science and Nutrition.
2004; 44(4): 223-237. [18] Han JH. Antimicrobial food packaging. Food Technology. 2000; 54(3): 56-65. [19] Le Loir Y, Baron F, Gautier M. Staphylococcus aureus and food poisoning. Genetics and Molecular Researches. 2003; 2(1): 63-
76. [20] Scannell AGM, Hill C, Ross RP, Marx S, Hartmeier W, Arendt EK. Development of bioactive food packaging materials using
immobilised bacteriocins Lacticin 3147 and Nisaplin®. International Journal of Food Microbiology. 2000; 60: 241-249. [21] Pranoto Y, Salokhe VM, Rakshit SK. Physical and antibacterial properties of alginate-based edible film incorporated with garlic
oil. Food Research International. 2005; 38: 267-272. [22] Seydim AC, Sarikus G. Antimicrobial activity of whey protein based edible films incorporated with oregano, rosemary and
garlic essential oils. Food Research International. 2006; 39(5): 639-644. [23] Millette M, Le Tien C, Smoragiewicz W, Lacroix M. Inhibition of Staphylococcus aureus on beef by nisin-containing modified
alginate films and beads. Food Control. 2007; 18: 878-884. [24] Fernandez-Saiz P, Lagaron JM, Hernandez-Muñoz P, Ocio MJ. Characterization of antimicrobial properties on the growth of S.
aureus of novel renewable blends of gliadins and chitosan of interest in food packaging and coating applications. International Journal of Food Microbiology. 2008; 124(1): 13-20.
[25] Li X, Xing Y, Jiang Y, Ding Y, Li W. Antimicrobial activities of ZnO powder-coated PVC film to inactivate food pathogens. International Journal of Food Science and Technology. 2009; 44: 2161-2168.
[26] Fernandez-Saiz P, Soler C, Lagaron JM, Ocio MJ. Effects of chitosan films on the growth of Listeria monocytogenes, Staphylococcus aureus and Salmonella spp. in laboratory media and in fish soup. International Journal of Food Microbiology. 2010; 137: 287-294.
[27] Vermeiren L, Devlieghere F, Debevere J. Effectiveness of some recent antimicrobial packaging concepts. Food Additives and Contaminants. 2002; 19: 163-171.
[28] Cooksey K. Effectiveness of antimicrobial food packaging materials. Food Additives and Contaminants. 2005; 22(10): 980-987. [29] Ozdemir M, Floros JD. Active food packaging technologies. Critical Reviews in Food Science and Nutrition. 2004; 44(3): 185-
193. [30] Ayana B, Turhan KN. Use of antimicrobial methylcellulose films to control Staphylococcus aureus during storage of Kasar
cheese. Packaging Technology and Science. 2009; 22(8): 461-469. [31] Buonocore GG, Conte A, Del Nobile MA. Use of a mathematical model to describe the barrier properties of edible films.
Journal of Food Science. 2005; 70(2): 142-147. [32] Outtara B, Simard RE, Piette G, Bégin A, Holley RA. Diffusion of acetic and propionic acids from chitosan-based antimicrobial
packaging films. Journal of Food Science. 2000; 65(5): 768-773. [33] Redl A, Gontard N, Guilbert S. Determination of sorbic acid diffusivity in edible wheat gluten and lipid based films. Journal of
Science against microbial pathogens: communicating current research and technological advances A. Méndez-Vilas (Ed.)_______________________________________________________________________________
