Polysaccharides Edible Films and Coatings: History
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Subjects: Materials Science, Coatings & Films
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Ewa Szpunar-Krok

There has been a significant increase in the development of edible films and coatings in recent times, and this is expected to have a significant impact on the quality of fruit and vegetables in the coming years. Consumers expect fresh fruit and vegetables free from pesticide residues, with high quality, nutritional value and an extended shelf life. The application of coatings and edible films to fruits and vegetables represents an environmentally friendly approach to an innovative solution to this problem. Coatings and edible films can act as ecological and biodegradable packaging. The coating strategy involves a combination of natural biopolymers and appropriate preservation methods. Numerous studies show that natural polysaccharides are well suited for use as packaging material for fresh fruit and vegetables and can often be an important alternative to synthetic compounds. Natural polymer materials are a good barrier to oxygen and carbon dioxide; however, they are characterised by excessive solubility in the water environment, water vapour permeability and low extensibility.

  • cellulose derivatives
  • chitosan and chitin
  • edible films and coatings

1. Definition and Methods of Applying Edible Films and Coatings

Edible films and coatings are defined as any thin material used to wrap or coat a product to extend its shelf life, which can be consumed with the product. They can improve the quality of, e.g., fruits and vegetables by protecting them from physical, chemical and microbiological changes such as moisture loss, enzymatic browning reactions and fat oxidation [1]. The thin layer formed by the edible materials that cover the fruit and vegetables is called the edible coating, while the layer previously shaped and placed on the raw material is called the edible film. The edible coatings are in liquid form and are applied to the raw material by immersing it in the solution. Edible films are shaped like a solid sheet and then applied by wrapping around the product [2][3][4].

For a long time, edible films and coatings have been used mainly as a one-component film or coating formulation. However, recently, a number of studies have been carried out on two-component and multi-component edible materials that provide better functional properties. In such a case, composite films or coatings are prepared by combining two or more binders to obtain structures with modified physical, mechanical and barrier properties that are superior to a single-component material. Therefore, various substances are used in film-forming preparations, such as plasticisers (glycerol, polyoxyethylene glycol, propylene glycol, sorbitol), cross-linkers, emulsifiers and enhancers (vegetable fatty acids) to improve or modify the basic functionality of the material [5][6][7][8]. However, introduced in excess, they may deteriorate mechanical properties and increase water vapour permeability (polypropylene glycol, sorbitol). Edible films and coatings have a number of advantages over synthetic coatings:

  • They act as a gas and moisture barrier that creates a modified atmosphere in the fruit, which, in turn, extends the shelf life and preserves the quality of fresh fruit and vegetables.
  • They act as a barrier against microorganisms and thus contribute to proper hygiene.
  • Several active ingredients such as anti-browning agents, dyes, aromas, nutrients and spices can be incorporated into the polymer matrix and consumed with the fruit, thus increasing safety and even nutritional and sensory properties.
  • They help to reduce synthetic packaging waste due to their biodegradable nature [2][4][9][10][11].

When creating edible films and coatings, the aim is to obtain durable, thin and homogeneous materials [12]. They can be produced by wet or dry methods. One of the wet production methods is to obtain edible films by removing the solvent used to prepare the film-forming solution. In this process, as a result of the physico-chemical intermolecular interaction, a continuous structure is formed and stabilised. The macromolecules of the film-forming solution are dissolved in a solvent (water, ethyl alcohol, acetic acid) and can be combined with other additives. The resulting film-forming solution is poured in a thin layer, dried and then removed from the surface [13]. The structure of the film is influenced by the drying conditions (temperature, relative humidity), the thickness of the film-forming solution layer and its composition [6][14][15]. Coatings are formed on the surface of fruit or vegetables from a liquid solution by dipping in a film-forming mixture, as well as by spraying or lubricating the surface [16]. This casting method is the most popular technique on a laboratory scale, although there are also some industrial-scale applications. The properties of the coating solution (density, viscosity, surface tension, food immersion speed) are important in this method [17]. The multilayer technique is used here, in which fruits and vegetables are electrolytically coated in layer by layer [13]. The coating process is carried out at an appropriate temperature in order to ensure the fluidity of the ingredients, create a specific thickness and even create coverage of the surface of fruits and vegetables. Cohesion of particles in the coating structure and adhesion between the coating and the surface of the plant product takes place during the coating process. The intensity of the cohesive forces directly affects the barrier and mechanical properties of the covering layer, which depend on the methods of formation [18].

Edible films and coatings can also be prepared by extrusion [19]. Extrusion is often preferred to casting as the method of fabricating films because the throughput of the process is faster and less energy is required for the removal of water [20]. The successful application of extrusion depends upon the main variables that should be controlled during the extrusion process. These variables include food polymer selection, liquid feed rates, screw configuration, screw speed, zone temperatures, product inlet/outlet pressures (and their differentials) as well as die configuration [21]. Although extrusion is a promising approach for the elaboration of edible films, there are a limited number of studies related to the use of this technology on pectin edible film elaboration. Probably, this is due to the behaviour and chemical interaction of food ingredients during extrusion, which is difficult to understand, along with many processing variables that need to be controlled [22][23]. Thus, Fishman et al. [20] used extrusion to prepare edible films from pectin/starch blends plasticised with glycerol. They used a twin-screw extruder with nine heating zones. As a result of the research, the authors found that the extruded edible films have a microstructure and thermodynamic mechanical properties comparable to those obtained by casting from a solution containing the same materials. Pectin films with various combinations of orange albedo and starch were produced under similar conditions [20]. In addition, Liu et al. [21] investigated optimal extrusion parameter conditions for the production of edible packaging films derived from food polymers. The authors indicated that the optimal conditions for the pectin films are temperatures in extruder zones 3 and zone 4 of 125 °C and 110 °C, respectively and a screw speed of 225 rpm. These conditions were determined on the basis of the evaluation of the physical and mechanical properties of the films, such as tensile strength, elongation, puncture resistance, colour, Young’s modulus, tear resistance, haze and thickness [21][23].

Films and coatings made from a single polymeric component are often brittle and breakable. To counteract this, a plasticiser, allowing the formation of an elastic structure, is introduced into the film-forming solution. The plasticiser enters the molecular chains of polymers and their physicochemical connections, thus increasing the coherence. The most commonly used plasticisers are polyols (glycerol, sorbitol, polyethylene glycol), sugars (glucose, honey) and lipids (monoglycerides, phospholipids). The appropriate selection of a plasticiser for films and coatings is important as it can significantly change the physicochemical properties of the coatings [13]. The methods and possibilities of coating fruits and vegetables depend on the type of product and its properties, storage and distribution conditions [24][25].

Scientists have investigated various polymers of biological origin to obtain a continuous structure of membranes or coatings. Polysaccharide hydrocolloids are the most common group of biopolymers used in the production of edible films and coatings. They can be obtained from sources such as plants, crustaceans and microorganisms. Cellulose derivatives, starches, alginates, pectins, chitosans, pullulan and carrageenans are the most popular polysaccharides used in the production of edible films and coatings [2]. However, these materials are hydrophilic in nature. Therefore, different types of oils and fats are incorporated into the hydrocolloid matrix to enhance their water vapour barrier properties. The most popular are waxes, triglycerides, acetylated monoglycerides, free fatty acids and vegetable oils [26].

2. Characteristics of Selected Polysaccharides

2.1. Seaweed Products—Alginate, Carrageenan and Agar

Alginate, carrageenan and agar are products obtained from species of marine algae belonging mainly to Phaeophyceae (brown algae) and Rhodophyceae (red algae) (Table 1).

Table 1. Occurrence of polysaccharides in marine algae [27].

Polysaccharide Phylum (Division)/Species of Seaweeds
Phaeophyceae Rhodophyceae
Alginates Macrocystis  
Laminaria
Ascophyllum
Ecklonia
Eisenia
Nereocystis
Sargassum
Carrageenans   Chondrus
Gigartina
Eucheuma
Hypnea
Iridaea
Kappaphycus
Gymnogongrus
Ahnfeltia
Furcellaria
Agar   Gelidium
Gracilaria
Pterocladia

Alginate, the sodium salt of alginic acid, is a brown seaweed product hat exhibits film-forming properties [6][28]. The colloidal properties of these biopolymers used as film or coating constituents are related to their chemical structure [29][30]. Alginate coating materials are made by the use of divalent cations such as Ca, Mg, Mn, Al and Fe, and it is used as a gelling agent [31][32]. Alginate consists of D-mannuronic acid (M) and L-guluronic acid (G) in various proportions, arrangements and molecular weights [27][29][33]. The physical properties of alginate gels largely depend, among others, on the ratio of guluronic acid to mannuronic acid residues, the successive order of these residues, the length of guluronic acid blocks and the total molecular weight of the polymer [33][34][35]. If the M/G ratio < 1, the gel indicates a large amount of guluronic acid, which is capable of forming strong bonds. If the M/G ratio > 1, the gel contains less guluronic acid, which may result in softer and more flexible structures [36]. On the other hand, alginic acid obtained from different species of brown algae may also be different, as it may contain three types of polymer segments and consist of (a) mainly D-mannuronic acid (M) units, (b) mainly L-guluronic acid units (G) or (c) alternating D-mannuronic acid and L-guluronic acid residues [27] (Table 2).

Table 2. Alginate composition of selected brown algae species.

Species Mannuronic Acid (M), wt % Guluronic Acid (G), wt % M:G Ratio References
Laminaria digitata 53–59 41–47 2.20–3.60 [27][37]
Laminaria hyperborea 30–38 62–70 0.45 [27][38][39]
Laminaria japonica 69 31 2.26 [40]
Ascophyllum nodosum 60–65 35–40 1.40–2.25 [27][37][38]
Ecklonia cava 62–67 33–38 1.60–2.00 [27][41]
Eisenia bicyclis 62 38 1.60 [27]
Macrocystis pyrifera 50–61 39–50 1.56 [28][37][40]
Durvillae antarctica 56 44 1.27 [42]
Sargassum ringgoldianum 64–70 30–36 1.80–2.30 [43]
Sargassum turbinaroides 49 52 0.94 [44]
Sargassum fluitans 34–54 46–66 0.52–1.18 [45]
Sargassum muticum 24 76 0.31 [46]
Sargassum polycystum 18 82 0.21 [46]
Nereocystis luetkeana 63 37 1.70 [47]
Pelvetia canaliculata 56–60 40–44 1.30–1.50 [48]
Undaria pinnatifida 60 40 1.50 [40]

The ability of alginate to irreversibly and immediately react with divalent and trivalent metal cations (Ca2+ and Ca3+) and to form water-insoluble polymers is used to form coatings of edible vegetables and fruit [49]. Alginates have good film-forming properties, giving a uniform, transparent, glossy look to films [2][31][50]. Films (membranes) made of alginates are impermeable to fats and oils, and like other hydrophilic polysaccharides, they are distinguished by high water vapour permeability [29][31][49]. Alginate has a unique colloidal property, which contains a stabilising, thickening, suspending film or coating, producing gel-forming and stabilising emulsion [32][49]. Alginate has some desirable properties, including reduction in shrinkage, moisture retention, colour and odour of food [31]. The alginate gel coating can protect vegetables and fruit from loss of turgor, as moisture is first lost from the edible coating before the protected parts of the edible plants are significantly dehydrated [50][51]. Alginate coatings provide a good barrier to oxygen, which contributes to the delay of lipid oxidation in vegetables and fruits and reduces their weight loss and the abundance of microflora on the surface [50][52][53]. They also contribute to delaying the ageing processes that take place in fruits and vegetables during their storage [54].

To extract the alginate, the seaweed is broken into pieces and mixed with a hot base solution (e.g., sodium carbonate). After about 2 h, the alginate dissolves as sodium alginate, forming a thick slurry that also has undissolved parts of the seaweed (glycellulose). The resulting solution is diluted with water and pressed through a filter cloth in a filter press along with an auxiliary filter, e.g., diatomaceous earth. The last step is the alginate precipitation from the filtered solution as alginic acid or calcium alginate [55][56]. Pretreatment of the seaweed with acid (prior to alkaline extraction) leads to a more efficient extraction, less coloured product and reduced viscosity loss during extraction due to the lower amount of phenolic compounds [57]. Due to the linear structure, alginate can form strong films and appropriate fibrous structures in the solid state; therefore, it is considered a good filmogenic material [36].

Carrageenans are natural hydrophilic polymers with a linear chain of partially sulphated galactans that have high membrane formation potential [26][58]. However, the production of edible films using carrageenans is not as popular as they are most commonly used as coatings [59]. They are extracted from the cell walls of red algae, most often from the species Chondrus crispus, Kappaphycus alvarezii and Eucheuma denticulatum [26][55] (Table 1). There are three types of carrageenan: kappa carrageenan (κ-carrageenan), iota carrageenan (ι-carrageenan) and lambda carrageenan (λ-carrageenan) [60]. The solubility of carrageenans in water depends on the content of ester sulphate and associated cations. Higher levels of ester sulphate mean a lower dissolution temperature. The presence of cations such as Na, K, Ca and Mg promotes cation-dependent aggregation between carrageenan helices. Depending on the concentration of the solution, the melting point of carrageenans is from 50 to 70 °C and the gelation temperature is from 30 to 50 °C [55][61]. Iota carrageenan forms elastic and clear gels with no syneresis in the presence of calcium salts [55]. Edible films based on iota carrageenan have good mechanical properties as they are emulsion stabilisers and can reduce oxygen transfer and limit surface dehydration and deterioration of fruit flavour [62]. Lambda carrageenan, on the other hand, does not form a gel but only forms highly viscous solutions; therefore, it cannot be used as an edible film [63]. Kappa carrageenan is one of the most common forms of carrageenan that can be used in food, and the gel made from it can be frozen and thawed. Kappa carrageenan has a double-helix conformation, and the linear helical parts can associate to form a three-dimensional gel in the presence of appropriate cations [64]. Kappa carrageenan creates strong and stiff gels with potassium salts and brittle gels with calcium salts. Kappa carrageenan gels are opaque but become clear when sugar is added [55]. These coatings, like others, obtained from seaweeds, effectively protect vegetables and fruit against loss of moisture and turgor, oxidation of compounds and ageing processes and, in combination with ascorbic acid, reduce the number of microorganisms [28].

Carrageenan was previously obtained by the method of extracting into an aqueous solution. After the filtrate containing the seaweed residues has been removed, the carrageenan should be recovered from the solution. However, this method is expensive, and nowadays carrageenan is obtained by a method in which the seaweed is washed to remove solid impurities and then treated with alkali to extract the carrageenan. After extraction, the diluted extracts (1–2% carrageenan) are filtered, concentrated and then precipitated with isopropanol until a fibrous coagulum is obtained. The coagulum is pressed to remove the solvent and washed. It is then dried and milled to the appropriate particle size [55].

Agar is a mixture of agarose (gelling fraction) and agaropectin (non-gelling fraction) that is obtained from red seaweed [65]. Agarose creates a supporting structure in the cell walls of red algae and is responsible for the gelling properties of agar, making it suitable for the formation of edible coatings of vegetables and fruit [66]. Agar is mainly made of galactose units with a regular alternation of L and D isomeric forms. Agar gel compared with kappa carrageenan has increased strength and a higher melting point, which is related to the lower content of anionic sulphates. Agar gel melts when heated to 85 °C and resets when cooled to 31–40 °C. However, the viscosity of agar in solution at 60 °C is lower than that of carrageenan. Agar has the ability to form strong, thermoreversible gels and is known for its hydrophilicity. Agar films and coatings are transparent, strong and stiff and insoluble in water under ambient conditions [61].

Agar is most often obtained by alkaline treatment followed by hot-water extraction. The alkaline treatment causes a chemical change in the agar (formation of 3,6-anhydro-galactopyranose), resulting in increased gel strength. Extraction with hot water (temperature around 100 °C) takes about 2–4 h, sometimes under pressure. The agar is dissolved in water, and the seaweed residues are removed by filtration. The agar is then recovered by alcohol precipitation [67].

2.2. Gums

Gum arabic (synonymous with acacia gum) is a polysaccharide obtained from the gummy exudate of the stems and branches of the species of the genus Acacia, most often of the species Acacia senegal (L.) Willd. var. senegal. Gum arabic can also be obtained from the following species: Acacia senegal var. karensis, Acacia seyal var. seyaland, Acacia seyal var. fistula, Acacia polyacantha, Acacia gerrardii, Acacia laeta, Acacia nilotica and Acacia fischeri [68]. Gum arabic is a complex, branched, anionic, hydrophilic heteropolysaccharide with a backbone of 1,3-linked β-galactopyranose units and side chains of 1,6-linked galactopyranose or arabinose units terminating in rhamnose or glucuronic acid or 4-O-methlglucuronic acid residues, which contains about 2% protein substance and is classified as an arabinogalactan–protein complex [68][69]. Depending on the source of the nodules, it may have a variable composition and different physical and chemical properties. Among the hydrocolloids, gum arabic is the least viscous and more soluble and has good emulsifying and film-forming properties [68][69][70][71][72]. Tara gum is obtained by ground endosperm of the seeds of the Cesalpinia spinosa tree. It consists of linear chains of (1 → 4)-β-D-mannopyranosyl residues having single side chain units of (1 → 6)-α-D-galactopyranosyl, in a ratio of 3:1. Galactomannans (including tara gum) have high viscosity, water-binding capacity and the ability to synergistically interact with other polymers. Tara gum, based on steric hindrance, possesses less galactose substitution compared with guar gum (other galactomannan), which is why it can make stronger films [73][74]. However, the edible film produced from it has relatively poor mechanical and water vapour barrier properties [75].

Guar gum is a hydrophilic carbohydrate biopolymer obtained from the seeds of Cyamopsis tetragonoloba. It consists of a linear long-chain molecule of β (1 → 4)-linked D-mannose residues with single linked α (1 → 6)-D-galactose, in which the ratio of mannose to galactose units varies from 1.5:1 to 1.8:1. Guar gum can form a homogeneous edible film that is almost completely soluble in water due to the large number of hydroxyl groups [76][77].

In turn, xanthan gum is an extracellular heteropolysaccharide produced by submerged aerobic fermentation of a pure Xanthomonas campestris culture [78][79]. Due to the ability to create highly viscous solutions at low concentrations and its biodegradability, it is used as an additive to starch-based films, improving some of its mechanical properties [80].

Basil seed gum (Ocimum basilicum L.) is an acidic (anionic) polysaccharide from which edible film can be made [81]. Basil seed gum consists of two main fractions of glucomannan (43%), with a glucose-to-mannose ratio of 10:2, and (1 → 4)-linked xylan (24.29%) and a minor fraction of glucan (2.31%). This gum can be used to produce films with good appearance and satisfactory mechanical properties. However, the improvement of the mechanical properties of this film, including extensibility and increased water solubility, can be obtained by adding glycerol [82].

Edible films and coatings made of polysaccharide gums form a semi-permeable barrier. This helps to maintain the nutritional value of fruit and vegetables, although weight loss may occur and the rate of respiration at the surface may be reduced. Edible coatings of this type not only improve the shelf life of fruits and vegetables and prevent their quality deterioration during storage but also protect them against pathogens [54][83].

2.3. Pectin

Pectin is a component of plant fibre and can be extracted from the plant cell walls. It is a complex anionic polysaccharide composed of β-1,4-linked α-D-galacturonic acid residues, where the uronic acid carboxyls are either fully (high methoxy pectin) or partially (low methoxy pectin) methyl-esterified [74][84]. Pectin-based films and coatings have excellent mechanical properties. Raw materials rich in pectin can be used as a potential natural plasticiser, improving the extensibility of edible films [85]. However, pectin-based films are poor moisture barriers and are therefore recommended for food with low moisture content [54]. Edible films produced from high methoxy pectin have more favourable features. In the process of plasticising pectin and starch with a high amylase content, strong and flexible edible and biodegradable films are obtained, which, after plasticising with glycerol, have good mechanical and oxygen barrier properties [84][86].

2.4. Cellulose Derivatives

The most common natural polymer in nature is cellulose. Cellulose consists of D-glucose units linked through β-1,4-glycosidic bonds. It is an almost linear polymer in which a highly crystalline structure is obtained due to the tight packing of anhydroglucose polymer chains. Cellulose is insoluble in water due to the large number of intramolecular hydrogen bonds. The water solubility of cellulose can be increased by treating with alkali so that its structure swells. For this, a reaction with methyl chloride, chloroacetic acid or propylene oxide is carried out to obtain methyl cellulose (MC), carboxymethyl cellulose (CMC), hydroxylpropyl cellulose (HPC) or hydroxypropyl methyl cellulose (HPMC) [84][87][88]. Methyl cellulose, hydroxypropyl methyl cellulose and hydroxylpropyl cellulose films have good film-forming properties and are also biodegradable, tasteless, odourless, flexible, of moderate strength, transparent, resistant to fats and oils as well as water-soluble and moderately permeable to moisture and gases (oxygen and carbon dioxide) [54][84][88]. Cellulose-based films and coatings, which have a large surface area and biopolymer structures, are able to retain most of water in the product and therefore exhibit anti-rancidity effect [89]. The water vapour permeability of these films is highly influenced by the hydrophobic:hydrophilic ratio of the film components [88]. Krochta et al. [90] showed that the characteristics of cellulose, including its gas and moisture barrier, were dependent on its molecular weight and that the higher the molecular weight, the better the properties. Edible films obtained with carboxymethyl cellulose are an effective barrier to the penetration of oxygen and carbon dioxide but have a poor water barrier [91]. Carboxymethyl cellulose-based edible films can be used as a suitable carrier for some probiotic strains and have significantly improved vapour permeability [92]. It has also been found that mixing carboxymethyl cellulose with other polymers, e.g., starch and chitosan, increases the inherent CMC shortcomings [54]. Methylcellulose is less hydrophilic and provides a better moisture barrier [84][88][93]. Moreover, it is widely used as a carrier of antibacterial agents in the production of edible films [90][92]. In turn, films based on hydroxypropyl methyl cellulose create a good gas barrier [94].

2.5. Starch

Starch accumulates primarily in the tubers, seeds and roots of plants. For industry, it is most often obtained from maize, wheat, edible cassava, potato, amaranth and quinoa [22]. Glucose polymerisation in the starch molecule results in the formation of two polysaccharide fractions, linear amylose and branched amylopectin, which together constitute about 98–99% of the dry weight of starch and are packed in concentric rings that form semi-crystalline and amorphous layers [95][96]. Both fractions consist of chains composed of α-D-glucopyranose residues linked by α-1,4-glycosidic bonds, and the chains, in turn, are linked by α-1,6-glycosidic bonds, thus creating branches in polymers. Amylose (20–30%) is a polymer with a linear structure, which greatly influences the amorphous form of starch granules [22][97]. Due to its linear structure, it tends to orient itself in parallel. This causes the formation of hydrogen bonds between hydroxyl groups, reducing the affinity of the polymer to water and, as a result, enabling the formation of membranes and gels [98]. Amylopectin (70–80%) has a highly branched structure, and its structure influences the peripheral crystal organisation of starch grains. Amylose and amylopectin are assembled naturally in granular forms, about 1–100 µm in size [22][99]. Moreover, the distribution of the unit chain length in the internal structure of amylose and amylopectin affects the thermal properties and the starch retrogradation profile [100]. The content of these components in starch determines the properties of the film, as edible films with higher amylose content have better film-forming properties, i.e., better mechanical strength, elongation and gas barrier properties [21]. To produce coatings with a higher amylose content, this polymer can be extracted from starch by selective leaching in hot water (50–70 °C) [101]. After cooling, starch forms a gel. During gelation, amylose and amylopectin undergo intra- and intermolecular cross-linking. As a result, macromolecular networks are formed [21][102].

Depending on the source of starch, the content of amylose and amylopectin varies significantly, as does the size of the granules (granules) and their shape (Table 3). These features affect the functionality, barrier, mechanical and sorption properties of the starch. On the other hand, the sorption properties of starch influence the composition of the coatings, the process of their formation, methods of applying to the product and application [103][104].

Table 3. Amylose and amylopectin content and granule size in various plant products.

Starch Source Amylose (%) Amylopectin (%) Average Granule Size (μm) References
Potato 21–30 70–79 36–100 [105][106][107]
Tapioca 17 83 14 [105]
Corn 19–26 74–81 13.3 [105][107]
Corn (waxy) 1 99 - [105][107]
Corn (high amylose) 50–85 15–50 9.8 [105]
Wheat 19–25 75–81 7–20 [105][107]
Rice 17 83 6–8 [106]
Pea (smooth) 33–50 50–67 2–40 [105]
Pea (wrinkled) 61–88 12–39 17–30 [105]
Amaranth 5–22 78–95 1–3 [106]

The use of starch as a component of edible coatings is possible due to its good barrier to gases (carbon dioxide, oxygen), adequate durability and cohesive strength. Moreover, starch films are odourless, tasteless, colourless and non-toxic [108][109]. However, a major limitation in the use of starch is the high water vapour permeability, which additionally adversely affects the mechanical properties of the coating [22][110]. Therefore, components with hydrophobic properties are often added in the film production phase, e.g., oleic acid and polyethylene glycol [111]. Starch does not dissolve in water, but with increasing water temperature, it strongly swells and dissolves in an alkaline environment. Starch films are produced by pouring starch dispersion on a smooth surface and then drying them [112]. When forming a starch film, first the starch granules are heated in excess water to prepare a viscous solution. However, aqueous solutions are usually unstable and tend to gel immediately after cooling due to the association of the polymer chains [113]. Therefore, at high temperatures and in excess of water, the starch undergoes a transformation known as gelatinisation, during which the starch granules change from the semi-crystalline phase to the amorphous state [114]. The loss of crystallinity occurs in two stages, the first of which involves swelling of the starch molecule at 60–70 °C [115] and the second, excessive swelling and solubilisation of the granule at temperatures above 90 °C, leading to a complete loss of structural integrity [116]. This process depends on the ratio of amylose to amylopectin, water content and temperature of the dispersion, which, in turn, influences the recrystallisation of starch during retrogradation. Once the chain interactions are complete, the weight of the gel is reduced by further evaporating the water until most of the free water is removed. During the retrogradation, the dissociated amylose and amylopectin chains in a gelatinised starch dispersion reunite to form more ordered structures. This process affects the permeability, solubility and mechanical properties of starch films [105][113].

2.6. Chitosan and Chitin

Chitin is a biopolymer that occurs naturally in the exoskeleton of crustaceans (shrimps, oysters, krill, crabs, squid, lobsters), the cell walls of filamentous fungi (Mucoraceae) and other biological materials such as arachnids and insects (bumblebees, crickets, bees, silkworm larval skin) [117][118][119]. Chitin consists mainly of poly(β-(1-4)-2-acetamido-D-glucose), and unlike structurally identical cellulose, the acetamide group here replaces the secondary hydroxyl on the second carbon atom of the repeating hexose unit (secondary hydroxyl on the second carbon atom of the hexose repeat unit) [120][121]. Chitosan is obtained by N-deacetylation of chitin in an alkaline environment [122]. It is a copolymer consisting of β-(1-4)-2-acetamido-D-glucose and β-(1-4)-2-amino-D-glucose units, with the latter usually exceeding 60% [120]. Chitosan is a non-toxic, non-allergenic, biodegradable, biocompatible and film-forming material with beneficial biological effects (antifungal, antibacterial, antitumour) [123][124][125]. Chitosan films are characterised by selective gas permeability (CO2 and O2) and good mechanical properties and are biodegradable, so they are not harmful to the environment [108].

Edible chitosan coating applied to the surface of fruit and vegetables reduces their respiration rate by regulating gas permeability [126]. However, due to high water vapour permeability, their use as edible films is limited, especially in humid environments [120][127]. Water vapour permeability is a parameter that ensures the organoleptic properties of stored vegetables and fruit, as well as the ability to fight against dehydration or rehydration of the film [128]. Another factor limiting the direct use of chitosan as an edible coating is its low water solubility, which results from its rigid crystal structure [125].

The antifungal and antimicrobial activity of chitosan results from its polycationic nature [129]. Its antibacterial activity is mediated by electrostatic forces between the protonated amino group (NH2) in chitosan and negative residues on the cell surface [130], while the greater number of protonated amino groups (NH2) present in chitosan is associated with an increase in deacetylation degrees (DD) [131]. According to Liu et al. [131] the bactericidal effect of chitosan is influenced by the electrostatic interaction between the NH3+ groups of chitosan and the phosphoryl groups of the phospholipid component of the cell membrane. Chitosan particles change the permeability of bacterial cell membranes, hindering the gas exchange of their cells with the environment, as well as lead to cell dysfunction through cell penetration or rupture of the cell membrane [132][133][134][135]. Chitosan forms edible coatings and films that protect vegetables and fruit against deterioration and microbial contamination, helping to maintain their quality and contributing to the extension of their shelf life [54][136].

About 30% of chitin is obtained from shrimp shells. In this process, proteins are removed by reaction with a weak NaOH solution (1–10%) at elevated temperatures (85–110 °C). After shredding the shells, a weak HCl solution (1–10%) is used at room temperature to remove CaCO3. The physicochemical properties of chitin are determined by the parameters of the extraction performed, e.g., temperature, concentration of the reagents used, their duration of action and the size of the shells. The basic parameters that characterise chitin are the degree of polymerisation, the degree of deacetylation and purity. When chitin becomes soluble in weak acids, it can be regarded as chitosan, which is formed during the treatment of chitin with a strong solution of NaOH (over 40%) at a temperature of 90–120 °C. In the deacetylation reaction, the acetyl groups (CH3CO−) are removed from the amino groups and soluble chitosan is formed. To obtain a soluble product, at least 65% of the acetyl groups should be removed from the chitin molecule. The degree of deacetylation depends on the duration of the reaction, temperature and concentration of NaOH solution [121]. The physicochemical characteristics of chitosan, e.g., molecular weight, polydispersity and purity, depend on the method used, the apparatus used and the source of the armour. Moreover, the total length of the polymer chain has a direct impact on its weight, and as the chain shortens, the water solubility of chitosan increases. The parameter determining the properties of chitosan is also the degree of chitin deacetylation, which, depending on the technological process, may be 70–100%. This parameter is important as it indicates the positive charge of the molecules when dissolved in a weak acid. An important parameter from the point of view of applications is also purity, which is determined by the content of such contaminants as proteins, dusts, insoluble compounds, bacteria and endotoxins. Contaminants block the active amino groups of chitosan, which reduces the effectiveness of its action. Therefore, it is important to precisely control the methods of its production.

The enzymatic process, as well as the chemical one, leads to the conversion of N-acetylglucosamine units into glucosamine units but is carried out under much milder conditions and is not associated with the simultaneous hydrolysis of the polymer chain. Chitin deacetylases take part in the enzymatic deacetylation process [137]. The enzymatic reaction can produce chitosan with a higher molecular weight and the desired degree of deacetylation compared to a polymer obtained by chemical processes [138]. Other enzymes involved in the conversion of chitin and chitosan include chitinases and chitosanases, which catalyse the hydrolysis of glycosidic bonds, but differ in their substrate specificity, hydrolysing chitin and chitosan bonds, respectively [139].

This entry is adapted from the peer-reviewed paper 10.3390/agronomy11050813

References

  1. Erkmen, O.; Barazi, A.O. General characteristics of edible flms. J. Food Biotechnol. Res. 2018, 2, 1–4.
  2. Mahajan, B.V.C.; Tandon, R.; Kapoor, S.; Sidhu, M.K. Natural Coatings for Shelf-Life Enhancement and Quality Maintenance of Fresh Fruits and Vegetables—A Review. J. Postharvest Technol. 2018, 6, 12–26.
  3. Okcu, Z.; Yavuz, Y.; Kerse, S. Edible films and coatings application in fruits and vegetables. Alinteri J. Agric. Sci. 2018, 33, 221–226.
  4. Oms-Oliu, G.; Rojas-Graü, M.A.; González, L.A.; Varela, P.; Soliva-Fortuny, R.; Hernando, M.I.H.; Munuera, I.P.; Fiszman, S.; Martín-Belloso, O. Recent approaches using chemical treatments to preserve quality of fresh-cut fruit: A review. Postharvest Biol. Technol. 2010, 57, 139–148.
  5. Radev, R.; Pashova, S. Application of Edible Films and Coatings for Fresh Fruit and Vegetables. Qual. Access Success 2020, 21, 108–112.
  6. Galus, S.; Kibar, E.A.A.; Gniewosz, M.; Kraśniewska, K. Novel Materials in the Preparation of Edible Films and Coatings—A Review. Coatings 2020, 10, 674.
  7. Shukla, P.; Bhise, S.; Thind, S.S. Role of biodegradable edible films and coatings in food industry. Acta Sci. Nutr. Health 2019, 3, 138–147.
  8. Salgado, P.R.; Ortiz, C.M.; Musso, Y.S.; Di Giorgio, L.; Mauri, A.N. Edible films and coatings containing bioactives. Curr. Opin. Food Sci. 2015, 5, 86–92.
  9. Tharantharn, R.N. Biodegradable films and composite coatings: Past, present and future. Trends Food Sci. Technol. 2003, 14, 71–78.
  10. Mohamed, A.Y.E.; Aboul, A.H.E.; Hassan, A.M. Utilisation of edible coating in extending the shelf life of minimally processed prickly pear. J. Appl. Sci. Res. 2013, 9, 1202–1208.
  11. Vaishali Sharma, H.P.; Shami, V.; Samsher; Chandhary, V.; Sunil; Kumar, M. Importance of edible coating on fruits and vegetables: A review. J. Pharmacogn. Phytochem. 2019, 8, 4104–4110.
  12. Fu, Z.; Wang, L.; Li, D.; Wei, Q.; Adhikari, B. Effects of high-pressure homogenization on the properties of starch-plasticizer dispersions and their films. Carbohydr. Polym. 2011, 86, 202–207.
  13. Basiak, E.; Galus, E.; Lenart, E. Starch films and coatings for food. Food Ind. 2012, 66, 28–30.
  14. Galus, S.; Lenart, A. Effect of coating on food quality. Technol. Prog. Food Process. 2019, 2, 106–114.
  15. Skurtys, O.; Acevedo, C.; Pedreschi, F.; Enrione, J.; Osorio, F.; Aguilera, J.M. Food Hydrocolloid Edible Films and Coatings; Nova Science Publishers Inc.: New York, NY, USA, 2010.
  16. Coma, V.; Martial–Gros, S.; Gareau, S.; Copinet, A.; Salin, F.; Dechamps, A. Edible antimicrobial films based on chitosan matrix. J. Food Sci. 2002, 67, 1162–1169.
  17. Cisneros-Zevallos, L.; Krochta, J.M. Dependence of coating thickness on viscosity of coating solution applied to fruits and vegetables by dipping method. J. Food Sci. 2003, 68, 503–510.
  18. Falguera, V.; Quintero, J.P.; Jimenez, A.; Munoz, A.; Ibarz, A. Edible films and coatings: Structures, active functions and trends in their use. Trends Food Sci. Technol. 2011, 22, 291–303.
  19. Ghanbarzadeh, B.; Almasi, H.; Entezami, A.A. Physical properties of edible modified starch/carboxymethyl cellulose films. Innov. Food Sci. Emerg. Tech. 2010, 11, 697–702.
  20. Fishman, M.L.; Coffin, D.R.; Onwulata, C.I.; Konstance, R.P. Extrusion of pectin and glycerol with various combinations of orange albedo and starch. Carbohydr. Polym. 2004, 57, 401–413.
  21. Liu, Z. Edible films and coatings from starches. In Innovations in Food Packaging; Han, J.H., Ed.; Academic Press: London, UK, 2005; pp. 318–337.
  22. Basiak, E.; Lenart, A. Starch coatings used in food packaging industry. Food Sci. Technol. Qual. 2013, 1, 21–31.
  23. Espitia, P.J.P.; Avena-Bustillos, R.J.; Du, W.-X.; Chiou, B.-S.; Williams, T.G.; Wood, D.; McHugh, T.H.; Soares, N.F.F. Physical and antibacterial properties of acai edible films formulated with thyme essential oil and apple skin polyphenols. J. Food Sci. 2014, 79, M903–M910.
  24. Hall, D.J. Edible coatings from lipids, waxes, and resins. In Edible Coatings and Films to Improve Food Quality, 2nd ed.; Baldwin, E.A., Hagenmaier, R., Bai, J., Eds.; CRC Press: Boca Raton, FL, USA, 2012; pp. 79–101.
  25. Araujo-Farro, P.C.; Podadera, G.; Sobral, P.J.A.; Menegalli, F.C. Development of films based on Quinna (Chenopodium quinoa, Willdenow) starch. Carbohyd. Polym. 2010, 81, 839–848.
  26. Tavassoli-Kafrani, E.; Shekarchizadeh, H.; Masoudpour-Behabadi, M. Development of edible films and coatings from alginates and carrageenans. Carbohydr. Polym. 2016, 137, 360–374.
  27. Glicksman, M. Utilization of seaweed hydrocolloids in the food industry. Hydrobiologia 1987, 151–152, 31–47.
  28. Lin, D.; Zhao, Y. Innovations in the Development and Application of Edible Coatings for Fresh and Minimally Processed Fruits and Vegetables. Compr. Rev. Food Sci. Food Saf. 2007, 6, 60–75.
  29. King, A.H. Brown seaweed extracts (alginates). In Food Hydrocolloids; Glicksman, E., Ed.; CRC Press: Boca Raton, FL, USA, 1983; pp. 115–188.
  30. Draget, K.I.; Smidsrod, O.; Skjak-Braek, G. Alginates from Algae. In Polysaccharides and Polyamides in the Food Industry. Properties, Production, and Patents; Steinbüchel, A., Rhee, S.K., Eds.; Wiley-Blackwell: Hoboken, NJ, USA, 2005; pp. 1–30.
  31. Borchard, W.; Kenning, A.; Kapp, A.; Mayer, C. Phase diagram of the system sodium alginate/water: A model for biofilms. Int. J. Boil. Macromol. 2005, 35, 247–256.
  32. Dhanapal, A.; Sasikala, P.; Rajamani, L.; Kavitha, V.; Yazhini, G.; Banu, M.S. Edible films from polysaccharides. Food Sci. Qual. Manag. 2012, 3, 9–18.
  33. Stabler, C.; Wilks, K.; Sambanis, A.; Constantinidis, I. The effects of alginate composition on encapsulated βTC3 cells. Biomaterials 2001, 22, 1301–1310.
  34. Klöck, G.; Pfeffermann, A.; Ryser, C.; Gröhn, P.; Kuttler, B.; Hahn, H.J.; Zimmermann, U.J.B. Biocompatibility of mannuronic acid-rich alginates. Biomaterials 1997, 18, 707–713.
  35. Draget, K.I.; Taylor, C. Chemical, physical and biological properties of alginates and their biomedical implications. Food Hydrocoll. 2011, 25, 251–256.
  36. Blanco-Pascual, N.; Montero, M.P.; Gómez-Guillén, M.C. Antioxidant film development from unrefined extracts of brown seaweeds Laminaria digitate and Ascophyllum nodosum. Food Hydrocoll. 2014, 37, 100–110.
  37. Haug, A.; Larsen, B. Quantitative determination of the uronic acid composition of alginates. Acta Chem. Scan. 1962, 16, 1908–1918.
  38. Grasdalen, H.; Larsen, B.; Smidsrød, O. A p.m.r. Study of the Composition and Sequence of Uronate Residues in Alginates. Carbohydr. Res. 1979, 68, 23–31.
  39. Penman, A.; Sanderson, G.R. A method for the determination of uronic acid sequence in alginates. Carbohydr. Res. 1972, 25, 273–282.
  40. Ji, M.; Wang, Y.; Xu, Z.; Guo, Y. Studies on the M:G ratios in alginate. Hydrobiologia 1984, 116/117, 554–556.
  41. Kim, D.S. Uronic Acid Composition, Block Structure and Some Related Properties of Alginic Acid. Master’s Thesis, Pusan Commercial (San Eup) University, Pusan, Korea, 1984.
  42. Draget, K.I.; Skjak, G.; Smidsrød, O. Alginic acid gels: The effect of alginate chemical composition and molecular weight. Carbohydr. Polym. 1994, 25, 31–38.
  43. Kim, D.S.; Park, Y.H. Uronic acid composition, block structure and some related properties of alginic acid. On alginic acid prepared from Sargassum ringgoldianum. Korean J. Fish. Aquat. Sci. 1985, 18, 29–36.
  44. Laroche, C.; Michaud, P. A novel alginate from the brown seaweed Sargassum turbinaroides (Sargassae). In Current Topics on Bioprocesses in Food Industry; Larroche, C., Pandey, A., Dussap, C.G., Eds.; Asiatech Publisher, Inc.: New Delhi, India, 2009; Volume 2, pp. 71–92.
  45. Davis, T.A.; Ramirez, M.; Mucci, A.; Larsen, B. Extraction, isolation and calcium binding of alginate from Sargassum spp. J. Appl. Physiol. 2004, 16, 275–284.
  46. Davis, T.A.; Llanes, F.; Volesky, B.; Mucci, A. Metal selectivity of Sargassum spp. and their alginates in relation to their a-L-guluronic acid content and conformation. Environ. Sci. Technol. 2003, 37, 261–267.
  47. Rosell, K.G.; Srivastava, L.M. Seasonal variation in the chemical constituents of the brown algae Macrocystis integrifolia and Nereocystis luetkeana. Can. J. Bot. 1984, 62, 2229–2236.
  48. Haug, A.; Larsen, B.; Smidsrød, O. Uronic acid sequence in alginate from different sources. Carbohydr. Res. 1974, 32, 217–225.
  49. Rhim, J.W. Physical and mechanical properties of water resistant sodium alginate films. LWT Food Sci. Technol. 2004, 37, 323–330.
  50. Kester, J.J.; Fennema, O.R. Edible films and coatings: A review. Food Technol. 1986, 40, 47–59.
  51. Han, J.H. Edible Films and Coatings: A Review. In Innovations in Food Packaging, 2nd ed.; Han, J.H., Ed.; Academic Press; Elsevier: Amsterdam, The Netherlands, 2014; Chapter 9; pp. 213–255.
  52. Bonilla, J.; Atarés, L.; Vargas, M.; Chiralt, A. Edible films and coatings to prevent the detrimental effect of oxygen on food quality: Possibilities and limitations. J. Food Eng. 2012, 110, 208–213.
  53. Conca, K.R.; Yang, T.C.S. Edible food barrier coatings. In Biodegradable Polymers and Packaging; Ching, C., Kaplan, D., Thomas, D., Eds.; Technomic Publishing Co. Inc.: Lancaster, PA, USA, 1993; pp. 357–369.
  54. Al-Tayyar, N.A.; Youssef, A.M.; Al-Hindi, R.R. Edible coatings and antimicrobial nanoemulsions for enhancing shelf life and reducing foodborne pathogens of fruits and vegetables: A review. Sustain. Mater. Technol. 2020, 26, e00215.
  55. McHugh, D.J. A guide to the seaweed industry. In FAO Fisheries Technical Paper 441; FAO: Rome, Italy, 2003.
  56. Hassan, B.; Chatha, S.A.S.; Hussain, A.I.; Zia, K.M. Recent advances on polysaccharides, lipids and protein based edible films and coatings: A review. Int. J. Biol. Macromol. 2018, 109, 1095–1107.
  57. McHugh, D.J. Production and utilization of products from commercial seaweeds. In FAO Fisheries Technical Paper 288; FAO: Rome, Italy, 1987.
  58. Osorio, F.A.; Molina, P.; Matiacevich, S.; Enrione, J.; Skurtys, O. Characteristics of hydroxy propyl methyl cellulose (HPMC) based edible film developed for blueberry coatings. Proc. Food Sci. 2011, 1, 287–293.
  59. Campos, C.A.; Gerschenson, L.N.; Flores, S.K. Development of edible films and coatings with antimicrobial activity. Food Bioproc. Tech. 2011, 4, 849–875.
  60. Cosenza, V.A.; Navarro, D.A.; Fissore, E.N.; Rojas, A.M.; Stortz, C.A. Chemical and rheological characterization of the carrageenans from Hypnea musciformis (Wulfen) Lamoroux. Carbohydr. Polym. 2014, 102, 780–789.
  61. Kapetanakou, A.E.; Manios, S.G.; Skandamis, P.N. Application of Edible Films and Coatings on Food. In Novel Food Preservation and Microbial Assessment Techniques; Boziaris, I.S., Ed.; CRC Press: Boca Raton, FL, USA; Taylor & Francis: Abingdon, UK, 2014; pp. 237–273.
  62. Hambleton, A.; Perpinan-Saiz, N.; Fabra, M.J.; Voilley, A.; Debeaufort, F. The schroeder paradox or how the state of water affects the moisture transfer through edible films. Food Chem. 2012, 132, 1671–1678.
  63. De Araújo, I.W.F.; de Sousa Oliveira Vanderlei, E.; Rodrigues, J.A.G.; Coura, C.O.; Quinderé, A.L.G.; Fontes, B.P.; Queiroz, I.N.L.; Jorge, R.J.B.; Bezerra, M.M.; de Silva, A.A.R.; et al. Effects of a sulfated polysaccharide isolated from the red seaweed Solieria filiformis on models of nociception and inflammation. Carbohydr. Polym. 2011, 86, 1207–1215.
  64. Lopez-Pena, C.L.; McClements, D.J. Optimizing delivery systems for cationic biopolymers: Competitive interactions of cationic polylysine with anionic kappa-carrageenan and pectin. Food Chem. 2014, 153, 9–14.
  65. Schmidt, É.C.; Dos Santos, R.; Horta, P.A.; Maraschin, M.; Bouzon, Z.L. Effects of UVB radiation on the agarophyte Gracilaria domingensis (Rhodophyta, Gracilariales): Changes in cell organization, growth and photosynthetic performance. Micron 2010, 41, 919–930.
  66. Rhim, J.W. Effect of clay contents on mechanical and water vapor barrier properties of agar-based nanocomposite films. Carbohydr. Polym. 2011, 86, 691–699.
  67. Li, L.; Ni, R.; Shao, Y.; Mao, S. Carrageenan and its applications in drug delivery. Carbohydr. Polym. 2014, 103, 1–11.
  68. Motlagh, S.; Ravines, P.; Karamallah, K.A.; Ma, Q. The analysis of Acacia gums using electrophoresis. Food Hydrocol. 2006, 20, 848–854.
  69. Murmu, S.B.; Mishra, H.N. Optimization of the arabic gum based edible coating formulations with sodium caseinate and tulsi extract for guava. LWT Food Sci. Technol. 2017, 80, 271–279.
  70. Elmanan, M.; Al-Assaf, S.; Philips, G.O.; Williams, P.A. Studies of Acacia exudates gums: Part IV. Interfacial rheology of Acacia senegal and Acacia seyal. Food Hydrocol. 2008, 22, 682–689.
  71. Ali, A.; Maqbool, M.; Ramachandran, S.; Alderson, P.G. Gum arabic as a novel edible coating for enhancing shelf-life and improving postharvest quality of tomato (Solanum lycopersicum L.) fruit. Postharvest Biol. Technol. 2010, 58, 42–47.
  72. Ali, A.; Maqbool, M.; Alderson, P.G.; Zahid, N. Effect of gum arabic as an edible coating on antioxidant capacity of tomato (Solanum lycopersicum L.) fruit during storage. Postharvest Biol. Technol. 2013, 76, 119–124.
  73. Antoniou, J.; Liu, F.; Majeed, H.; Qazi, H.J.; Zhong, F. Physicochemical and thermomechanical characterization of tara gum edible films: Effect of polyols as plasticizers. Carbohydr. Polym. 2014, 111, 359–365.
  74. Chen, Y.; Xu, L.; Wang, Y.; Chen, Z.; Zhang, M.; Chen, X. Characterization and functional properties of a pectin/tara gum based edible film with ellagitannins from the unripe fruits of Rubus chingii Hu. Food Chem. 2020, 325, 126964.
  75. Antoniou, J.; Liu, F.; Majeed, H.; Zhong, F. Characterization of tara gum edible films incorporated with bulk chitosan and chitosan nanoparticles: A comparative study. Food Hydrocol. 2015, 44, 309–319.
  76. Aydogdu, A.; Radkea, C.J.; Bezci, S.; Kirtil, E. Characterization of curcumin incorporated guar gum/orange oil antimicrobial emulsion films. Int. J. Biol. Macromol. 2020, 148, 110–120.
  77. Banegas, R.S.; Zornio, C.F.; Borges, A.M.G.; Porto, L.C.; Soldi, V. Preparation, Characterization and Properties of Films Obtained from Cross-linked Guar Gum. Polimeros 2013, 23, 182–188.
  78. Arismendi, C.; Chillo, S.; Conte, A.; Del Nobile, M.A.; Flores, S.; Gerschenson, L.N. Optimization of physical properties of xanthan gum/tapioca starch edible matrices containing potassium sorbate and evaluation of its antimicrobial effectiveness. LWT Food Sci. Technol. 2013, 53, 290–296.
  79. Ge, L.; Li, X.; Zhang, R.; Yang, R.; Ye, X.; Li, D.; Mu, D. Development and characterization of dialdehyde xanthan gum crosslinked gelatin based edible films incorporated with amino-functionalized montmorillonite. Food Hydrocol. 2015, 51, 129–135.
  80. Guo, J.M.; Ge, L.M.; Li, X.Y.; Mu, C.D.; Li, D.F. Periodate oxidation of xanthan gum and its crosslinking effects on gelatin-based edible films. Food Hydrocol. 2014, 39, 243–250.
  81. Gahruie, H.H.; Mostaghimi, M.; Ghiasi, F.; Tavakoli, S.; Naseri, M.; Hosseini, S.M.H. The effects of fatty acids chain length on the techno-functional properties of basil seed gum-based edible films. Inter. J. Biol. Macromol. 2020, 160, 245–251.
  82. Khazaei, N.; Esmaiili, M.; Djomeh, Z.E.; Ghasemlou, M.; Jouki, M. Characterization of new biodegradable edible film made from basil seed (Ocimum basilicum L.) gum. Carbohydr. Polym. 2014, 102, 199–206.
  83. Otoni, C.G.; Avena-Bustillos, R.J.; Azeredo, H.M.C.; Lorevice, M.V.; Moura, M.R.; Mattoso, L.H.C.; McHugh, T.H. Recent advances on edible films based on fruits and vegetables—A review. Compr. Rev. Food Sci. Food Saf. 2017, 16, 1151–1169.
  84. Dhall, R.K. Advances in Edible Coatings for Fresh Fruits and Vegetables: A Review. Crit. Rev. Food Sci. Nutr. 2013, 53, 435–450.
  85. Jahromi, M.; Niakousari, M.; Golmakani, M.T.; Mohammadifar, M.A. Physicochemical and structural characterization of sodium caseinate based film-forming solutions and edible films as affected by high methoxyl pectin. Int. J. Biol. Macromol. 2020, 165, 1949–1959.
  86. Coffin, D.R.; Fishman, M.L. Physical and mechanical properties of highly plasticized pectin/starch films. J. Appl. Polym. Sci. 1994, 54, 1311–1320.
  87. Bourtoom, T. Edible films and coatings: Characteristics and properties. Int. Food Res. J. 2008, 15, 237–248.
  88. Šuput, D.Z.; Lazić, V.L.; Popović, S.Z.; Hromiš, N.M. Edible films and coatings—Sources, properties and application. Food Feed Res. 2015, 42, 11–22.
  89. Kalkan, S.; Otağ, M.R.; Engin, M.S. Physicochemical and bioactive properties of edible methylcellulose films containing Rheum ribes L. extract. Food Chem. 2020, 307, 125524.
  90. Krochta, J.M.; Mulder-Johnson, C. Edible and biodegradable polymer films challenges and opportunities. Food Tech. 1997, 52, 661–674.
  91. Li, H.; Shi, H.; He, Y.; Fei, X.; Peng, L. Preparation and characterization of carboxymethyl cellulose-based composite films reinforced by cellulose nanocrystals derived from pea hull waste for food packaging applications. Int. J. Biol. Macromol. 2020, 164, 4104–4112.
  92. Ebrahimi, B.; Mohammadi, R.; Rouhi, M.; Mortazavian, A.M.; Shojaee-Aliabadi, S.; Koushki, M.R. Survival of probiotic bacteria in carboxymethyl cellulose based edible film and assessment of quality parameters. LWT Food Sci. Technol. 2018, 87, 54–60.
  93. Zillo, R.R.; da Silva, P.P.M.; de Oliveira, J.; da Glória, E.M.; Spoto, M.H.F. Carboxymethylcellulose coating associated with essential oil can increase papaya shelf life. Sci. Hortic. 2018, 239, 70–77.
  94. Choi, W.S.; Singh, S.; Lee, Y.S. Characterization of edible film containing essential oils in hydroxypropyl methylcellulose and its effect on quality attributes of ‘Formosa’ plum (Prunus salicina L.). LWT Food Sci. Technol. 2016, 70, 213–222.
  95. Wani, A.A.; Singh, P.; Shah, M.A.; Schweiggert-Weisz, U.; Gul, K.; Wani, I.A. Rice starch diversity: Effects on structural, Morphological, Thermal, and Physicochemical Properties—A Review. Compr. Rev. Food Sci. Food Saf. 2012, 11, 417–436.
  96. Copeland, L.; Blazek, J.; Salman, H.; Tang, M.C. Form and functionality of starch. Food Hydrocoll. 2009, 23, 1527–1534.
  97. Tester, R.F.; Karkalas, J.; Qi, X. Starch—composition, fine structure and architecture. J. Cereal Sci. 2004, 39, 151–165.
  98. Guimarãesac, I.C.; Cristina dos Reis, K.; Galvão, E.; Menezes, E.G.T.; Rodrigues, A.C.; Ferreira da Silva, T.; Nolasco de Oliveira, I.R.; Vilas Boas, E.V.B. Cellulose microfibrillated suspension of carrots obtained by mechanical defibrillation and their application in edible starch films. Ind. Crop. Prod. 2016, 89, 285–294.
  99. Zhu, F. Modifications of starch by electric field based techniques. Trends Food Sci. Technol. 2018, 75, 158–169.
  100. Matignon, A.; Tecante, A. Starch retrogradation: From starch components to cereal products. Food Hydrocol. 2017, 68, 43–52.
  101. Torres, F.G.; Troncoso, O.P.; Torres, C.; Diaz, D.A.; Amaya, E. Biodegradability and mechanical properties of starch films from Andean crops. Int. J. Biol. Macromol. 2011, 48, 603–606.
  102. Zavareze, E.R.; Pinto, V.Z.; Klein, B.; Halal, S.L.M.E.; Elias, M.C.; Prentice-Hernández, C.; Dias, A.R.G. Development of oxidised and heat-moisture treated potato starch film. Food Chem. 2012, 132, 344–350.
  103. Li, Y.; Shoemaker, C.F.; Ma, J.; Shen, X.; Zhong, F. Paste viscosity of rice starches of different amylose content and carboxymethylcellulose formed by dry heating and the physical properties of their films. Food Chem. 2008, 109, 616–623.
  104. Mali, S.; Grossmann, M.V.E.; Garcia, M.A.; Martino, M.N.; Zaritzky, N.E. Effects of controlled storage on thermal, mechanical and barrier properties of plasticized films from different starch sources. J. Food Eng. 2006, 75, 453–460.
  105. Thakur, R.; Pristijono, P.; Scarlett, C.J.; Bowyer, M.; Singh, S.P.; Vuong, Q.V. Starch-based films: Major factors affecting their properties. Int. J. Biol. Macromol. 2019, 132, 1079–1089.
  106. Januszewska-Jóźwiak, K.; Synowiecki, J. Characteristics and suitability of amaranth components in food biotechnology. Biotechnologia 2008, 3, 89–102.
  107. Pietrzyk, S.; Fortuna, T. Impact of starch type and its oxidation conditions on retrogradation properties. Food Sci. Technol. Qual. 2005, 2, 23–32.
  108. Pająk, P.; Fortuna, T.; Przetaczek-Rożnowska, I. Protein and polysaccharide-based edible packagings: Profile and applications. Food Sci. Technol. Qual. 2013, 2, 5–18.
  109. Shah, U.; Naqash, F.; Gani, A.; Masoodi, F.A. Art and Science behind Modified Starch Edible Films and Coatings: A Review. Compr. Rev. Food Sci. Food Saf. 2016, 15, 568–580.
  110. Rodríguez, M.; Osés, J.; Ziani, K.; Maté, J.I. Combined effect of plasticizers and surfactants on the physical properties of starch based edible films. Food Res. Int. 2006, 39, 840–846.
  111. Ryu, S.Y.; Rhim, J.W.; Roh, H.J.; Kim, S.S. Preparation and physical properties of zein-coated high amylose corn starch film. LWT Food Sci. Technol. 2002, 35, 680–686.
  112. de Moraes, J.O.; Scheibe, A.S.; Sereno, A.; Laurindo, J.B. Scale-up of the production of cassava starch based films using tape-casting. J. Food Eng. 2013, 119, 800–808.
  113. Wang, S.; Li, C.; Copeland, L.; Niu, Q.; Wang, S. Starch retrogradation: A comprehensive review. Compr. Rev. Food Sci. Food Saf. 2015, 14, 568–585.
  114. Ratnayake, W.S.; Jackson, D.S. Starch gelatinization. Adv. Food Nutr. Res. 2009, 55, 221–268.
  115. Shanks, R.; Kong, I. Thermoplastic Starch. In Thermoplastic Elastomers; El-Sonbati, A.Z., Ed.; InTechOpen: London, UK, 2012; Available online: (accessed on 11 February 2021).
  116. Ivanič, F.; Jochec-Mošková, D.; Janigová, I.; Chodák, I. Physical properties of starch plasticized by a mixture of plasticizers. Eur. Polym. J. 2017, 93, 843–849.
  117. Zhang, M.; Haga, A.; Sekiguchi, H.; Hirano, S. Structure of insect chitin isolated from beetle larva cuticle and silkworm (Bombyx mori) pupa exuvia. Int. J. Biol. Macromol. 2000, 27, 99–105.
  118. Majtan, J.; Bilikora, K.; Markowiec, O.; Grof, J.; Kogan, G.; Simuth, J. Isolation and characterization of chitin from bumblebee (Bombus terrestris). Int. J. Biol. Macromol. 2007, 40, 237–241.
  119. Sajomsang, W.; Gonil, P. Preparation and characterization of α-chitin from cicada sloughs. Mater. Sci. Eng. C 2010, 30, 357–363.
  120. Elsabee, M.Z.; Abdou, E.S. Chitosan based edible films and coatings: A review. Mater. Sci. Eng. C 2013, 33, 1819–1841.
  121. Tyliszczak, B.; Bialik-Wąs, K.; Drabczyk, A.; Kudłacik, S.; Sobczak-Kupiec, A. Animal-derived chitosans. Characteristics, comparison, application. Przem. Chem. 2016, 95, 2059–2062.
  122. Butola, B.S. Recent advances in chitosan polysaccharide and its derivatives in antimicrobial modification of textile materials. Int. J. Biol. Macromol. 2019, 121, 905–912.
  123. Domard, A.; Domard, M. Chitosan: Structure–properties relationship and biomedical applications. In Polymeric Biomaterials; Severian, D., Ed.; Marcel Decker Incorporated: New York, NY, USA, 2001; pp. 187–212.
  124. Youssef, A.M.; Abdel-Aziz, M.S.; El-Sayed, S.M. Chitosan nanocomposite films based on Ag-NP and Au-NP biosynthesis by Bacillus subtilis as packaging materials. Int. J. Biol. Macromol. 2014, 69, 185–191.
  125. Chaudhary, S.; Kumar, S.; Kumar, V.; Sharma, R. Chitosan nanoemulsions as advanced edible coatings for fruits and vegetables: Composition, fabrication and developments in last decade. Int. J. Biol. Macromol. 2020, 152, 154–170.
  126. Hajji, S.; Younes, I.; Affes, S.; Boufi, S.; Nasri, M. Optimization of the formulation of chitosan edible coatings supplemented with carotenoproteins and their use for extending strawberries postharvest life. Food Hydrocoll. 2018, 83, 375–392.
  127. Candir, E.; Ozdemir, A.E.; Aksoy, M.C. Effects of chitosan coating and modified atmosphere packaging on postharvest quality and bioactive compounds of pomegranate fruit cv. ‘Hicaznar’. Sci. Hortic. 2018, 235, 235–243.
  128. Elsabee, M.Z. Chitosan-Based Edible Films. In Polysaccharides; Ramawat, K., Mérillon, J.M., Eds.; Springer: Cham, Switzerland, 2014; pp. 1–37.
  129. Kim, K.W.; Thomas, R.L.; Lee, C.; Park, H.J. Antimicrobial Activity of Native Chitosan, Degraded Chitosan, and O-Carboxymethylated Chitosan. J. Food Prot. 2003, 66, 1495–1498.
  130. Tsai, G.; Su, W.; Chen, H.; Pan, C. Antimicrobial activity of shrimp chitin and chitosan from different treatments and applications of fish preservation. Fish. Sci. 2002, 68, 70–177.
  131. Liu, H.; Du, Y.; Wang, X.; Sun, L. Chitosan kills bacteria through cell membrane damage. Int. J. Food Microbiol. 2004, 95, 147–155.
  132. Goy, R.C.; Morais, S.T.; Assis, O.B. Evaluation of the antimicrobial activity of chitosan and its quaternized derivative on E. coli and S. aureus growth. Rev. Bras. Farm. 2016, 26, 122–127.
  133. Singh, R.; Shitiz, K.; Singh, A. Chitin and chitosan: Biopolymers for wound management. Int. Wound J. 2017, 14, 1276–1289.
  134. Pavinatto, A.V.; de Almeida Mattos, A.C.G.; Malpass, M.H.; Okura, D.T.; Balogh, R.C.; Sanfelice, A. Coating with chitosan-based edible films for mechanical/biological protection of strawberries. Int. J. Biol. Macromol. 2020, 151, 1004–1011.
  135. Sharaf, O.M.; Al-Gamal, M.S.; Ibrahim, G.A.; Dabiza, N.M.; Salem, S.S.; El-Ssayad, M.F.; Youssef, A.M. Evaluation and characterization of some protective culture metabolites in free and nano-chitosan-loaded forms against common contaminants of Egyptian cheese. Carbohydr. Polym. 2019, 223, 115094.
  136. Bakshia, P.S.; Selvakumara, D.; Kadirvelub, K.; Kumara, N.S. Chitosan as an environment friendly biomaterial—A review on recent modifications and applications. Int. J. Biol. Macromol. 2019, 150, 1072–1083.
  137. Kaczmarek, M.B.; Struszczyk-Swita, K.; Li, X.; Szczęsna-Antczak, M.; Daroch, M. Enzymatic Modifications of Chitin, Chitosan, and Chitooligosaccharides. Front. Bioeng. Biotechnol. 2019, 7, 243.
  138. Tsigos, I.; Martinou, A.; Kafetzopoulos, D.; Bouriotis, V. Chitin deacetylases: New, versatile tools in biotechnology. Trends Biotechnol. 2000, 18, 305–312.
  139. Jaworska, M.M. Kinetics of enzymatic deacetylation of chitosan. Cellulose 2012, 19, 363–369.
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