Figures 3–5 show the spectra of new samples and three composted variants (Figure 3 sample no. 5, with the addition of TDPA; Figure 4—sample no. 7, with the addition of d2w®; Figure 5—the sample described by the manufacturer as a bag for organic waste no. 12), that did not decompose. Figures showing the FTIR spectra of the remaining samples are included in the supplementary materials. The spectra were obtained during the X-raying of new samples (NEW), composted without irradiation (A), exposed to UV radiation for 20 h and composted (B) and UV irradiated for 50 h and composted (C). The Table of Characteristic IR Absorptions, published by the University of Colorado in Boulder, Department of Chemistry and Biochemistry, was used to analyse the spectra [39,40].

Figure 3 shows the spectra obtained during the X-ray of sample no. 5 (new and three composted variants). The bands visible at the wave numbers of approximately 2915 and 2847 cm−1 correspond to the stretching vibrations of the methylene C–H group. Initially, C–H stretching occurs, and then there is a marked drop in the peak, corresponding to bond cleavage. This applies to all of the samples, including the new ones. However, in the case of the coloured part of the material, the peaks are most intense for sample C, i.e., irradiated for 50 h and then composted. Further changes occur only at approximately 1472 cm−1 (occurrence of C–H bending vibrations). The presence of the bands in the wave number range 1320–1000 cm−1 is small. The growth of the bands around 1116 and 1111 cm−1 confirms the appearance of oxygen groups C–O in carboxylic and ester bonds. The bands

at approximately 874, 730 and 718 cm−1 correspond to the C–Cl stretching vibrations. During visual inspection, no significant changes were noted for sample no. 5 after the composting process. *Materials* **2021**, *14*, x FOR PEER REVIEW 8 of 15

**Figure 3.** Sample no. 5 spectra: (**a**) spectra generated by the white part of the sample material, (**b**) spectra generated by the coloured part of the sample material. **Figure 3.** Sample no. 5 spectra: (**a**) spectra generated by the white part of the sample material, (**b**) spectra generated by the coloured part of the sample material.

In the case of sample no. 7 (Figure 4), the first significant changes appeared already in the range of wave numbers amounting to approximately 3394 cm−1, which proves the occurrence of stretching vibrations of the amino group N–H. These changes are most intense for the coloured part of sample B, i.e., irradiated for 20 h and then composted. One can also see very intense peaks at approximately 2915, 2909, 2844 and 2847 cm−1, which correspond to the vibrations of the C–H methylene group. Initially, C–H stretching occurs, followed by bond cleavage. This applies to all the samples, including the new ones. Both in the white and coloured parts of the material, peaks in this range are the most intense for trial C, i.e., irradiated for 50 h and then composted. Further changes occur in the spectra of coloured materials at the wave numbers of approximately 1619 cm−1, which corresponds to the appearance of bending vibrations of the N–H amino group. The peaks In the case of sample no. 7 (Figure 4), the first significant changes appeared already in the range of wave numbers amounting to approximately 3394 cm−1 , which proves the occurrence of stretching vibrations of the amino group N–H. These changes are most intense for the coloured part of sample B, i.e., irradiated for 20 h and then composted. One can also see very intense peaks at approximately 2915, 2909, 2844 and 2847 cm−1 , which correspond to the vibrations of the C–H methylene group. Initially, C–H stretching occurs, followed by bond cleavage. This applies to all the samples, including the new ones. Both in the white and coloured parts of the material, peaks in this range are the most intense for trial C, i.e., irradiated for 50 h and then composted. Further changes occur in the spectra of coloured materials at the wave numbers of approximately 1619 cm−1 , which corresponds to the appearance of bending vibrations of the N–H amino group. The peaks

at wave numbers around 1461 and 1462 cm−1 (for white and coloured material) are the reappearance of C–H bonds (bending vibrations). The next changes concern the wave numbers in the range from approximately 1116 and 1107 cm−1 and confirm the appearance of C–O oxygen groups of carboxylic and ester bonds. The bands at approximately 718 cm−1 are C–Cl stretching vibrations. Finally, in the coloured part of sample no. 7B, a very intense band appears at a wave number of approximately 599 cm−1 . According to the authors of the tables [39,40] these are stretching vibrations of C–Br bonds, alkyl halides, which are very stable compounds. The presented changes can be related to the visual assessment. After the composting process, we observed clear changes in the coloured part of sample no. 7. The sample material was also less flexible. at wave numbers around 1461 and 1462 cm−1 (for white and coloured material) are the reappearance of C–H bonds (bending vibrations). The next changes concern the wave numbers in the range from approximately 1116 and 1107 cm−1 and confirm the appearance of C–O oxygen groups of carboxylic and ester bonds. The bands at approximately 718 cm−1 are C–Cl stretching vibrations. Finally, in the coloured part of sample no. 7B, a very intense band appears at a wave number of approximately 599 cm−1. According to the authors of the tables [39,40] these are stretching vibrations of C–Br bonds, alkyl halides, which are very stable compounds. The presented changes can be related to the visual assessment. After the composting process, we observed clear changes in the coloured part of sample no. 7. The sample material was also less flexible.

*Materials* **2021**, *14*, x FOR PEER REVIEW 9 of 15

**Figure 4.** Sample no. 7 spectra: (**a**) spectra generated by the white part of the sample material, (**b**) spectra generated by the coloured part of the sample material. **Figure 4.** Sample no. 7 spectra: (**a**) spectra generated by the white part of the sample material, (**b**) spectra generated by the coloured part of the sample material.

In the case of sample no. 12 (Figure 5) the first changes appear in all samples in the range of wave numbers amounting to approximately 3236 cm−1 , and this is the occurrence of stretching vibrations of the amino group N–H. The bands visible at the wave numbers of approximately 2914 and 2847 cm−1 correspond to the vibrations of the methylene C–H

group (stretching and bond cleavage). Changes at approximately 1639 cm−1 correspond to the appearance of bending vibrations of the N–H amino group. Further changes occur at approximately 1460 cm−1 (occurrence of C–H bending vibrations). The growth of the bands around 1101 and 1032 cm−1 is the appearance of oxygen groups C–O in carboxylic and ester bonds. The bands at approximately 874 and 718 cm−1 are C–Cl stretching vibrations. Sample no. 12 was made entirely of a brown-coloured material which did not change its structure after the composting process, and the colour changes were minor. The material was still flexible and durable. The producers suggested choosing this type of bag for collecting organic waste (as indicated by the description on the packaging), but even a visual assessment aroused doubts regarding its purpose. The structure, appearance and flexibility of the bag were almost identical to those made of polyethylene. of approximately 2914 and 2847 cm−1 correspond to the vibrations of the methylene C–H group (stretching and bond cleavage). Changes at approximately 1639 cm−1 correspond to the appearance of bending vibrations of the N–H amino group. Further changes occur at approximately 1460 cm−1 (occurrence of C–H bending vibrations). The growth of the bands around 1101 and 1032 cm−1 is the appearance of oxygen groups C–O in carboxylic and ester bonds. The bands at approximately 874 and 718 cm−1 are C–Cl stretching vibrations. Sample no. 12 was made entirely of a brown-coloured material which did not change its structure after the composting process, and the colour changes were minor. The material was still flexible and durable. The producers suggested choosing this type of bag for collecting organic waste (as indicated by the description on the packaging), but even a visual assessment aroused doubts regarding its purpose. The structure, appearance and flexibility of the bag were almost identical to those made of polyethylene.

In the case of sample no. 12 (Figure 5) the first changes appear in all samples in the range of wave numbers amounting to approximately 3236 cm−1, and this is the occurrence of stretching vibrations of the amino group N–H. The bands visible at the wave numbers

*Materials* **2021**, *14*, x FOR PEER REVIEW 10 of 15

**Figure 5.** Spectra of sample no. 12 (the entire sample was made up of brown material). **Figure 5.** Spectra of sample no. 12 (the entire sample was made up of brown material).

Figure 6 shows the carbonyl index values for the samples after composting. In most cases, the indexes achieved higher values for samples that were exposed to UV radiation for 20 h before composting (B). This does not confirm the results of the spectra analysis, which showed some Figure 6 shows the carbonyl index values for the samples after composting. In most cases, the indexes achieved higher values for samples that were exposed to UV radiation for 20 h before composting (B).

changes (indicating the ongoing degradation process) in some of the samples irradiated for 50 h. The changes were not clear, they indicated that the decomposition of the samples was starting, even though the composting process was finished. The use of hierarchical cluster analysis (supplementary materials, Figure S6) con-This does not confirm the results of the spectra analysis, which showed some changes (indicating the ongoing degradation process) in some of the samples irradiated for 50 h. The changes were not clear, they indicated that the decomposition of the samples was starting, even though the composting process was finished.

firmed the existence of similar relationships between the spectra of the tested samples. Samples that were completely decomposed during composting (1, 2, 3, 11) ended up in the first main cluster (A), along with the color part of sample no. 6. According to the manufacturer's declaration, 85% of it was made from sugar cane and there were signs of a biodegradation process. Samples 1, 2, 3 and 11 could be made of materials containing similar components. The remaining samples were included in the second cluster (B). It includes all the samples that were not decomposed in the composting process (except for The use of hierarchical cluster analysis (supplementary materials, Figure S6) confirmed the existence of similar relationships between the spectra of the tested samples. Samples that were completely decomposed during composting (1, 2, 3, 11) ended up in the first main cluster (A), along with the color part of sample no. 6. According to the manufacturer's declaration, 85% of it was made from sugar cane and there were signs of a biodegradation process. Samples 1, 2, 3 and 11 could be made of materials containing similar components. The remaining samples were included in the second cluster (B). It includes all the samples that were not decomposed in the composting process (except for the coloured part of sample no. 6). This cluster is divided into two smaller ones: the first one (B0 ) is dominated by coloured parts of samples (4, 10, 5, 8) and samples of uniform colour (9 and 12). It also includes the white parts of samples 4 and 5. Most of the mentioned samples showed an increase in the CI value after 20 h irradiation compared to the samples composted without irradiation. The second cluster (B") has the most white samples (6, 8, 7, 10), and the coloured part of sample number 7 belongs to this.

**Figure 6.** Values of the carbonyl indexes for samples after composting: A—composted without irradiation, B—exposed to UV radiation for 20 h and composted, C—irradiated for 50 h and composted. **Figure 6.** Values of the carbonyl indexes for samples after composting: A—composted without irradiation, B—exposed to UV radiation for 20 h and composted, C—irradiated for 50 h and composted.

#### **4. Discussion 4. Discussion**

Synthetic polymer packaging is readily available, flexible and resistant to many factors. However, they are not renewable, non-biodegradable and pose a threat to the environment. Packages made of natural raw materials (biodegradable and ecological) are usually very light, thin and have low durability. They are not resistant to factors such as water, air and high temperature. It is very difficult to obtain a material that is both biodegradable, environmentally safe and durable [41]. Work on finding the golden mean in this area is ongoing. New types of material are appearing that may replace those previously used. There are many different packages available on the market for consumers described by producers as biodegradable and oxo-biodegradable. The information on these packages is not always accurate. Our research has shown that not all packaging decomposes, despite the fact that the research conditions corresponded to real-world conditions. Another scenario, e.g., longer exposure to UV rays, additional mechanical actions, heating, are not possible or available under real composting conditions. Synthetic polymer packaging is readily available, flexible and resistant to many factors. However, they are not renewable, non-biodegradable and pose a threat to the environment. Packages made of natural raw materials (biodegradable and ecological) are usually very light, thin and have low durability. They are not resistant to factors such as water, air and high temperature. It is very difficult to obtain a material that is both biodegradable, environmentally safe and durable [41]. Work on finding the golden mean in this area is ongoing. New types of material are appearing that may replace those previously used. There are many different packages available on the market for consumers described by producers as biodegradable and oxo-biodegradable. The information on these packages is not always accurate. Our research has shown that not all packaging decomposes, despite the fact that the research conditions corresponded to real-world conditions. Another scenario, e.g., longer exposure to UV rays, additional mechanical actions, heating, are not possible or available under real composting conditions.

the coloured part of sample no. 6). This cluster is divided into two smaller ones: the first one (B′) is dominated by coloured parts of samples (4, 10, 5, 8) and samples of uniform colour (9 and 12). It also includes the white parts of samples 4 and 5. Most of the mentioned samples showed an increase in the CI value after 20 h irradiation compared to the samples composted without irradiation. The second cluster (B″) has the most white

samples (6, 8, 7, 10), and the coloured part of sample number 7 belongs to this.

All samples analysed with the use of FTIR spectroscopy were characterised by differences in the obtained spectra. The samples that did not decompose in the composting process were characterised by intense bands in the 3000–2800 cm−1 range, which correspond to the stretching and cleavage of the C–H group bonds. These bands are characteristic, for example, for polypropylene [42], they did not occur in samples 1, 2, 3 and 11, which completely decomposed in the composting process. All of the samples also had new peaks below 1500 cm−1, known as the dactyloscopic range. Bands of stretching vibrations of single bonds appear (for example, C–H, C–Cl), corresponding to deformation vibrations, as well as groups of C–O oxygen carboxylic and ester bonds. The observed changes in the spectra indicate the occurrence of the oxidation and degradation process of the materials used in the samples. The spectra of samples 4, 7, 8 and 9, belonging to the films described as oxo-biodegradable, are similar to those presented by Benitez et al., who studied the degradation of polyethylenes with pro-oxidants added. The authors also found that these types of material oxidize the components and degrade the material [43,44]. Also in the case of sample no. 6, some changes appeared, indicating the occurrence of the sample decomposition process (especially the coloured part). In the case of All samples analysed with the use of FTIR spectroscopy were characterised by differences in the obtained spectra. The samples that did not decompose in the composting process were characterised by intense bands in the 3000–2800 cm−1 range, which correspond to the stretching and cleavage of the C–H group bonds. These bands are characteristic, for example, for polypropylene [42], they did not occur in samples 1, 2, 3 and 11, which completely decomposed in the composting process. All of the samples also had new peaks below 1500 cm−1 , known as the dactyloscopic range. Bands of stretching vibrations of single bonds appear (for example, C–H, C–Cl), corresponding to deformation vibrations, as well as groups of C–O oxygen carboxylic and ester bonds. The observed changes in the spectra indicate the occurrence of the oxidation and degradation process of the materials used in the samples. The spectra of samples 4, 7, 8 and 9, belonging to the films described as oxo-biodegradable, are similar to those presented by Benitez et al., who studied thedegradation of polyethylenes with pro-oxidants added. The authors also found that thesetypes of material oxidize the components and degrade the material [43,44]. Also in thecase of sample no. 6, some changes appeared, indicating the occurrence of the sample decomposition process (especially the coloured part). In the case of samples no. 7 and 8, the analysis of the spectra revealed the presence of compounds from the group of alkyl halides (C–Br bonds). These compounds are used in the chemical industry and can pose a threat to the environment. Bromine is used in the production of dyes, and its compounds are persistent and stable in the environment. Some of the dyes can also be carcinogenic and mutagenic [45]. Perhaps due to the intense colour of the samples (prints, inscriptions), this compound appeared in them.

Registered changes were found for both white and coloured material. However, they prove abiotic degradation, i.e., the process of initial oxidation of the sample material. This is the stage before proper biodegradation with the use of microorganisms [46–48]. It should be noted that the results relate to samples that have already been treated in the composting process. Therefore, they cannot be considered biodegradable, nor can they be considered to degrade in the environment or in compost. In this case, the analysis of spectra obtained using FTIR spectroscopy cannot be treated as the only confirmation of biodegradability. They should be supplemented with other types of test. The carbonyl index allows us to determine the amount of carbonyl compounds formed during the photo-oxidation process. However, it has been shown that a significant part of the oxidation product is omitted, because it evaporates into the atmosphere. Therefore, this indicator cannot be considered as a reliable probe to measure the extent of oxidation and does not reflect the total degradation of the mechanical properties of the polymer [49]. Moreover, the conditions under which biodegradability was tested are important. If these were laboratory conditions, they also do not have to demonstrate the ability to be completely biodegradable, because such conditions are different from real ones and this information may mislead the consumer.

The conducted tests were aimed at assessing the degree and nature of changes occurring during the decomposition of selected packaging and confirming that at various stages of material biodegradation, harmful microplastics and substances contained in plastics may be released into the compost. The processing of plastics together with organic waste causes the pollution to enter the compost, which can then be used as fertiliser [50].

Analysis of the spectra also showed that earlier irradiation of samples did not bring the expected results. According to some authors, earlier exposure of plastics to UV radiation accelerates biodegradation [51] or increases growth of carbonyl indicators [52]. The results show only partial degradation of the samples. In samples no. 5, 6 and 7 there were greater differences between the spectra of samples irradiated for 50 h and the others. Visual inspection confirmed that the structure of some samples had been disturbed. An earlier publication presented the content of selected components in samples before and after the composting process [36]. Analyses have shown that elements such as zinc, copper, chromium and lead can get into the compost. During the decomposition of biodegradable and oxo-biodegradable plastics, microplastic particles may also form, which accumulate in the environment. This was also confirmed by other authors [53,54]. Microplastics are a potential threat to flora and fauna [55], and they affect the functions of soil and microbial communities [56]. The biodegradation of packaging used as waste bags or for storing food is very important, because it concerns many elements of the environment. Errors resulting from a too rash assessment of biodegradability potential may pose a threat to the environment and even to the health and life of animals.

#### **5. Conclusions**

The analysis of the results of the conducted research allowed for the formulation of the following conclusions:

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/ 10.3390/ma14216449/s1, Figure S1: Sample no. 4 spectra: (a) spectra generated by the white part of the sample material, (b) spectra generated by the coloured part of the sample material, Figure S2: Sample no. 6 spectra: (a) spectra generated by the white part of the sample material, (b) spectra generated by the coloured part of the sample material, Figure S3: Sample no. 8 spectra: (a) spectra generated by the white part of the sample material, (b) spectra generated by the coloured part of the sample material, Figure S4: Spectra of sample no. 9 (the entire sample was made up of white material), Figure S5: Sample no. 10 spectra: (a) spectra generated by the white part of the sample material, (b) spectra generated by the coloured part of the sample material, Figure S6: Dendrogram showing the analyzed new samples. Agglomeration was carried out using the Ward method.

**Author Contributions:** Conceptualization, F.M. and A.S.-P.; methodology, F.M. and A.S.-P.; software, A.S.-P.; validation, F.M. and A.S.-P., formal analysis, F.M. and A.S.-P.; investigation, F.M.; resources, F.M.; data curation, A.S.-P.; writing—original draft preparation, F.M. and A.S.-P.; writing—review and editing, F.M. and A.S.-P.; visualization, F.M. and A.S.-P.; supervision, A.S.-P.; project administration, F.M.; funding acquisition, F.M. All authors have read and agreed to the published version of the manuscript.

**Funding:** The research was carried out as part of a targeted subsidy for research and development of young scientists and PhD students at the Faculty of Environmental Engineering and Geodesy of the Wrocław University of Environmental and Life Sciences—contract no. B030/0111/18.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data presented in this study are available on request from the corresponding author.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**

*Article*

## **Whey Protein Concentrate**/**Isolate Biofunctional Films Modified with Melanin from Watermelon (***Citrullus lanatus***) Seeds**

**Łukasz Łopusiewicz 1,\* , Emilia Drozłowska 1 , Paulina Trocer 1 , Mateusz Kostek 1 , Mariusz Sliwi ´nski ´ 2 , Marta H. F. Henriques 3,4 , Artur Bartkowiak 1 and Peter Sobolewski 5**

Received: 6 August 2020; Accepted: 1 September 2020; Published: 2 September 2020

**Abstract:** Valorization of food industry waste and plant residues represents an attractive path towards obtaining biodegradable materials and achieving "zero waste" goals. Here, melanin was isolated from watermelon (*Citrullus lanatus*) seeds and used as a modifier for whey protein concentrate and isolate films (WPC and WPI) at two concentrations (0.1% and 0.5%). The modification with melanin enhanced the ultraviolet (UV) blocking, water vapor barrier, swelling, and mechanical properties of the WPC/WPI films, in addition to affecting the apparent color. The modified WPC/WPI films also exhibited high antioxidant activity, but no cytotoxicity. Overall, the effects were melanin concentration-dependent. Thus, melanin from watermelon seeds can be used as a functional modifier to develop bioactive biopolymer films with good potential to be exploited in food packaging and biomedical applications.

**Keywords:** melanin; watermelon seeds; whey protein; bioactive films; plant residues

#### **1. Introduction**

The food market represents a large part of the global economy and is growing every year. Hand-in-hand, this economic sector is now also responsible for approx. 1.3 billion tons of waste per annum [1]. This waste, from fruit, vegetable, and food, includes waste generated during all aspects of food production: cleaning, processing, cooking, and packaging. However, some of these waste products and/or by-products can be important sources of bioactive compounds, such as phenolic compounds, dietary fiber, polysaccharides, vitamins, carotenoids, pigments, and oils [2]. These compounds can be potentially used in the development of novel food products (food additives and functional foods) or food packaging materials. This is an attractive path towards waste valorization in line with current market trends connected with "zero waste" goals and the so-called circular economy [3,4]. Therefore, continuing research into both the characterization and utilization of compounds obtained from food-industry waste/by-products is important, because it may offer a path towards improved sustainability of the food industry. This could significantly mitigate environmental problems associated with this industry, as well as have a positive impact from the point of view of climate change [1,2,5].

Watermelon (*Citrullus lanatus*, clade: Rosids, order: Cucurbitales, family: *Cucurbitaceae*) is a very popular fruit, with the flesh both consumed, as well as processed into juice and juice concentrates, due to water content approaching 92% of total weight. However, watermelon seeds, which constitute about 1 to 4% of total fruit weight, are not routinely eaten with the pulp [6–9]. At the same time, these seeds do have economic value, particularly in countries where cultivation is increasing. They can be used to prepare snacks or be milled into flour and used in sauces. Watermelon seeds are reported to be a rich source of proteins, vitamins B and E, minerals (such as magnesium, potassium, phosphorous, sodium, iron, zinc, manganese and copper), polyunsaturated fatty acids such as omega-6 (linoleic acid), and monounsaturated fatty acids, such as omega-9 (oleic acid). They also consist of saturated fatty acids, such as palmitic acid and stearic acid, and were found to be rich in γ-sitosterol, β-sitosterol, and lupeol [6–9]. Further, they are a promising source of useful compounds with potential biofunctional properties such as polyphenols, saponins, alkaloids and flavonoids [10,11]. However, despite these applications, watermelon seeds are still typically discarded, with only the fruit being eaten [10,12].

Biodegradable edible films are defined as a thin layer of material, that can be consumed. They are typically used to extend the shelf life and/or to improve the quality of foods. For example, they can be used to act as barriers to mass transfer, carriers of specific ingredients, or for the improvement of mechanical/handling characteristics of the product [13–15]. Growing consumer demand for high-quality foods, along with increasing environmental concern regarding the disposal of non-renewable food packaging materials, has led to a great deal of interest in the development of novel, biodegradable edible films/coatings [1,15,16]. However, such films, which are typically composed of biopolymers, can be also used in biomedical applications i.e., as wound dressings [17]. Further, the functional properties of such biopolymer films can be improved by adding different biofunctional compounds (e.g., antioxidant and/or antimicrobial properties). In this fashion, one can obtain biodegradable, bioactive materials with properties suitable for a range of diverse applications, while reducing the use of synthetic chemical additives that may have negative on human health or the environment [1,15,16].

At present, the packaging industry is dominated by synthetic polymers (plastics), because they are very cheap and possess good mechanical and physical properties. The annual plastic production is estimated to be approx. 300 million tons, of which 40% is used in packaging. However, this wide use of synthetic packaging materials has caused serious concerns, due to their high environmental impact [1,18]. Synthetic packaging polymers are petroleum-based and thus non-renewable, while at the same time being typically non-biodegradable. As a result, packaging accounts for large amounts of waste materials and pollution in the environment [1,13,19]. As a result, there is a pressing need to develop new, more eco-friendly packaging materials. In this context, biopolymers are very promising, because, compared to petroleum-based synthetic plastics, they are derived from a biological origin, making them renewable, biodegradable, and non-toxic or biocompatible [1,14]. A wide range of carbohydrates, proteins, and lipids—all derived from renewable sources—are being investigated as biodegradable alternatives, to improve sustainability and recyclability [1,13,20,21]. In particular, protein-based films are promising, due to better mechanical attributes, barrier characteristics, and nutritional-promoting properties, as compared to polysaccharide and lipid-based materials [20,22]. Gradually, bio-sourced materials are likely to replace the commonly utilized petroleum-based polymers, as environmental and sustainability externalities become increasingly accounted for in their cost [23].

Among protein-based edible films, whey protein (WP) films have received increased interest, because they possess interesting sensorial, optical, and mechanical barrier properties [24,25]. Whey is a protein-rich, major by-product of the cheese manufacturing industry [1,24,26]. In fact, this industry generates large volumes of fluid whey, that need to be properly disposed of, in order to avoid potential environmental problems [25]. Thus, whey protein-based edible films and coatings are not only

value-added products, but also offer a potential solution to the disposal problem [1]. Heat-denatured whey proteins, with the addition of a plasticizer, yield transparent, bland, and flexible films with very good resistance to oxygen, aroma, and lipid transfer at low humidity [15,23]. However, the hydrophilic nature of the proteins enables interactions with water, which leads to a reduction in the moisture barrier properties [25,27]. In addition to applications in edible films, whey protein concentrates and isolates (WPC, WPI) also have the potential to be used in the biomedical field, for example forming hydrogels as bioactive carriers or by leveraging their antioxidative properties [28–31].

Melanins are black and brown biopigments, consisting of high molecular weight heterogeneous polymers derived from the oxidation of monophenols and the subsequent polymerization of intermediate *o*-diphenols and their resulting quinones [32]. The molecular structure of melanins includes multiple different reactive functional groups (−OH, −NH, and −COOH) [21]. They can be obtained and have been characterized by a variety of natural sources, including animals, plants, bacteria, and fungi [10,11,33,34]. Importantly, melanins are multifunctional and biologically-active, natural macromolecules and can be characterized as antioxidant, radioprotective, thermo-regulative, chemoprotective, antitumor, antiviral, antimicrobial, immunostimulating and/or anti-inflammatory [10,32,34–36]. Potentially, melanins could be used to impart some of these important attributes to polymers. In the case of biopolymers, this could enhance performance, as well as sustainability credentials. Further, melanins could enable a wide range of applications, for example by facilitating cross-linking during polymerization, providing antioxidant or antimicrobial activity, altering light scattering ability, or improving other biological properties of the polymers [18,19,37]. Importantly, melanins, like biopolymers, are obtained from renewable resources and are non-toxic; these two features make their use "greener" than many existing commercial additives [35]. Importantly, large-scale production of melanins by microorganisms digesting food waste, as well as by sustainable extraction from natural plant-residues (e.g., watermelon seeds) have been demonstrated [10,11,34]. In fact, melanin from watermelon seeds has been shown to have antioxidant and UV-barrier properties [10]. However, compared to their potential, the use of melanins remains under-explored. The relatively few examples typically involve their blending/use with polymers (as chemical modifiers or nanofillers) to modify films and coatings, for example: gelatin [38,39], poly(lactic acid) [18], alginate [21], agar [19], carrageenan [13], cellulose [22], chitosan [14], poly(vinyl alcohol) [40], polypropylene/poly(butylene adipate-co-terephthalate) [37], polyhydroxybutyrate [41] and, ethylene-vinyl acetate copolymer [35].

In this study, our aim was to investigate the effect of adding melanin obtained from watermelon seed on the properties of whey protein concentrate/isolate (WPC/WPI) films. To the best of our knowledge, no reports have been published on the modification of WPC or WPI films with natural melanin to improve the functionality of the materials. We used UV-Vis and IR spectroscopy to examine the chemical composition of films after melanin addition. Additionally, we also assessed the influence of melanin on the color, hydrodynamic, and optical properties of the films. Finally, in order to evaluate the potential (bio) functionality of the obtained materials, we evaluated their mechanical, barrier, and antioxidant properties and screened for any potential cytotoxicity in vitro.

#### **2. Materials and Methods**

#### *2.1. Materials and Reagents*

Whey protein concentrate (WPC, 85% protein content) and whey protein isolate (WPI, 90% protein content) manufactured from sweet cheese whey using cross-flow membrane filtration were purchased from Volac International Ltd. (Hertfordshire, UK). Calcium chloride, hydrogen peroxide, disodium phosphate, monosodium phosphate, 2,2-diphenyl-1-picrylhydrazyl (DPPH), 2,20 -azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS), potassium persulphate, potassium ferricyanide, trichloroacetic acid, ferric chloride, iron sulphate, tris(hydroxymethyl)aminomethane, pyrogallol, ortophenantroline, L929 murine fibroblasts, Dulbecco's Modified Eagle Medium (DMEM), fetal bovine serum (FBS), resazurin, l-glutamine, penicillin, streptomycin, and all other cell culture

reagents were purchased from Sigma Aldrich (Darmstad, Germany). Glycerol, ammonia water, hydrochloric acid, sodium hydroxide, chloroform, ethyl acetate, ethanol and methanol were supplied from Chempur (Piekary Sl ˛askie, Poland). Cell culture plasticware was purchased from VWR ´ International (Radnor, PA, USA). All chemicals were of analytical grade.

#### *2.2. Isolation, Purification and Preparation of Melanin Powder*

Fresh Crimson Sweet watermelons (*Citrullus lanatus*) were purchased at a local market (Szczecin, Poland). Melanin isolation and purification were performed as described previously [10]. Briefly, watermelon seeds were first manually removed, then rinsed three times with distilled water, and finally dried at room temperature. Then, melanin was extracted by soaking 5 g of seeds in 50 mL of 1 M NaOH on an orbital shaker (150 rpm, 50 ◦C, 24 h), followed by centrifugation (6000× *g* rpm, 10 min) to remove plant tissue. Next, in order to precipitate the melanin, 1 M HCl was added to the alkaline mixture until the pH was 2.0, followed by centrifugation (6000× *g* rpm, 10 min). Then, the resultant pellet was first hydrolyzed in 6 M HCl (90 ◦C, 2 h), centrifuged (6000× *g* rpm, 10 min), and washed with distilled water five times to remove acid. After this procedure, in order to remove lipids and other residues, the pellet was washed with chloroform, ethyl acetate, and ethanol three times. Thus obtained, the purified melanin was dried and ground to a fine powder in a mortar.

#### *2.3. Preparation of WPC and WPI Films*

WPC/WPI-based films were prepared based on the methodology of Catarino et al. with minor modifications [24]. Briefly, film-forming solutions with a protein concentration of 10% (*w*/*w*) WPC or WPI were prepared in distilled water, at room temperature under continuous stirring. Once completely dissolved, ammonia water was added to adjust the pH to 8.0. Next, melanin was added to obtain concentrations of 0.1% and 0.5% (*w*/*w*) and stirred (250 rpm) for 1 h, until the melanin was completely dissolved. This mixture was then heated for 10 min in a water bath at 90 ◦C, until a uniform appearance was observed. Next, the mixture was cooled to room temperature and 5% (*w*/*w*) of glycerol (on a film-forming solution basis) was added, followed by homogenization. As reference materials, neat WPC/WPI films, without melanin addition, were also produced following the same procedure. All film samples were prepared in 10 repetitions. The film-forming solutions were cast on square (120 mm × 120 mm) polystyrene plates and dried at 40 ◦C for 48 h. Then, the dry films were carefully peeled off of the plates and conditioned at 25 ◦C and 50% RH in the clean room, prior to any tests.

#### *2.4. Determination of Moisture Content, Water Solubility and Swelling Ratio*

The moisture content (MC), water solubility (WS), and swelling ratio (SR) of obtained films were analyzed following the methodology of Roy et al. [14]. In brief, MC was determined as the weight change of the films after drying at 105 ◦C for 24 h. To determine the water solubility (WS), film specimens (2.5 cm × 2.5 cm) were first dried at 60 ◦C overnight and then weighed. The dried films were then dipped in 30 mL of distilled water for 24 h with occasional shaking at 25 ◦C, then carefully removed with a tweezer, and dried at 105 ◦C for 24 h, and finally re-weighed. The WS of the films was then calculated using the following formula:

$$\text{WS} \left( \% \right) = \frac{\mathcal{W}\_1 - \mathcal{W}\_2}{\mathcal{W}\_1} \times 100 \tag{1}$$

where *W*1 is the initial and *W*2 is the final weight of the films, respectively.

To determine the SR of the films, pre-weighed samples were submerged in 30 mL of distilled water for 1 h. Then, surface water was carefully removed using filter paper and the samples were re-weighed. The following formula was used to calculate SR:

$$\text{SR (\%)} = \frac{W\_2 - W\_1}{W\_1} \times 100\tag{2}$$

where *W*1 is initial and *W*2 is the final weight of the films, respectively.

#### *2.5. Thickness, Mechanical, and Thermal Properties of WPC*/*WPI Films*

The thickness of all obtained films was measured using a hand-held micrometer (Dial Thickness Gauge 7301, Mitoyuto Corporation, Kangagawa, Japan) with an accuracy of 0.001 mm. Each film was measured in five random points and the results were averaged.

The mechanical properties of the obtained films were tested using a Zwick/Roell 2,5 Z universal testing machine (Ulm, Germany). Static tensile testing was carried out to assess tensile strength and elongation at break (The gap between tensile clamps was 25 mm and crosshead speed was 100 mm/min).

Differential scanning calorimetry (DSC) measurements to assess thermal properties were carried out using a DSC calorimeter (DSC 3, Mettler-Toledo LLC, Columbus, OH, USA) over a temperature range from 30 to 300 at ϕ = 10◦ /min and under nitrogen flow (50 mL/min), performing two heating and one cooling scans.

#### *2.6. The Water Vapour Transmission Rate (WVTR) of the Films*

A gravimetric method was used to determine the Water Vapour Transmission Rate (WVTR) of the obtained films, as described previously [18]. This method relies on the sorption of humidity by calcium chloride. Briefly, 9 g of dry CaCl2 was placed inside a container and sealed with 8.9 cm2 samples of each film. Over the course of four days, the containers were weighed daily and the increase in mass indicated that water vapor passed through the films. For each film type, 10 film samples were tested, and average values for each day were calculated and used to express WVTR in g/(m2 × day).

#### *2.7. The Water Contact Angle (WCA)*

The water contact angle of all obtained films was measured using a Haas µL goniometer (Pozna ´n, Poland). Briefly, for each film, a microsyringe was used to deposit a drop of water on the surface. Three drops were analyzed and the contact angles were averaged.

#### *2.8. Spectral Analysis*

The UV-Vis spectra (300–700 nm) of the film samples were measured using a UV-Vis Thermo Scientific Evolution 220 spectrophotometer (Waltham, MA, USA).

Infrared spectroscopy was used in order to assess the chemical composition of obtained films, as described previously [18]. Briefly, 4 cm2 squares of each film were placed directly on the ray-exposing stage of the ATR accessory of a Perkin Elmer Spectrum 100 FT-IR spectrometer (Waltham, MA, USA) operating in ATR mode. Spectra (64 scans) were recorded over a wavenumber range of 650–4000 cm−1 , at a resolution of 4 cm−1 . For analysis, spectra were baseline corrected and normalized using SPECTRUM software [18].

#### *2.9. Color Analysis*

The effect of melanin on the color of the films was measured using a colorimeter (CR-5, Konica Minolta, Tokyo, Japan). For each film type, five samples were analyzed, by making three measurements on both sides of each sample. The results (mean ± standard deviation) were expressed as L\* (lightness), a\* (red to green), and b\* (yellow to blue). Additionally, ∆E (color difference) and YI (yellowness index), compared to unmodified WPC/WPI films, were also calculated as follows:

$$\Delta \mathbf{E} = \left[ (\mathbf{L}\_{\text{standard}} - \mathbf{L}\_{\text{sample}})^2 + (\mathbf{a}\_{\text{standard}} - \mathbf{a}\_{\text{sample}})^2 + (\mathbf{b}\_{\text{standard}} - \mathbf{b}\_{\text{sample}}) \right]^{0.5} \tag{3}$$

$$\text{YI} = \text{142.86b} \text{\*} \text{L}^{-1} \tag{4}$$

#### *2.10. Antioxidant Potential of the Films*

#### 2.10.1. Reducing Power

The reducing power of the films was determined based on the previously described methodology [42] with our own modification. Briefly, film samples (100 mg) were placed in 1.25 mL of phosphate buffer (0.2 M, pH 6.6), followed by the addition of 1.25 mL of 1% potassium ferricyanide solution. Samples were then incubated for 20 min at 50 ◦C followed by the addition of 1.25 mL of trichloroacetic acid. Next, the test tubes were centrifuged at 3000× *g* rpm for 10 min and 1.25 mL of obtained supernatant was diluted with 1.25 mL of deionized water. Finally, 0.25 mL of 0.1% ferric chloride solution was added and the absorbance was measured at 700 nm.

#### 2.10.2. Free Radical Scavenging Activity

The free radical scavenging activity of WPC/WPI films was assessed towards ABTS, DPPH, superoxide (O2 −), and hydroxyl (·OH) radicals. ABTS and DPPH tests were performed according to Bishai et al. with a slight modification [43]. For the ABTS test, 5 mL of 7 mM ABTS was mixed with 5 mL of 2.45 mM potassium persulfate to obtain the radical 2,20 -azino-bis(3-ethylbenzothiazoline)-6-sulphonic acid (ABTS+). After 16 h of incubation at room temperature protected from light, the solution was diluted to an absorbance maximum of 1.00 at 734 nm using water. To 25 mL of this ABTS+ solution, samples of each film (100 mg) were added and incubated up to 1 h at room temperature. As a control, tubes of ABTS+ solution were incubated under identical conditions, but without films. Finally, absorbance was measured and ABTS scavenging was calculated using the equation:

$$\text{Free radical saving activity (\%)} = \frac{A\_{\text{sample}} - A\_{\text{control}}}{A\_{\text{sample}}} \times 100,\tag{5}$$

where *Asample* is the absorbance of ABTS+ solution with the film sample and *Acontrol* is the absorbance of ABTS+ solution without sample.

To determine DPPH radical scavenging activity, 100 mg of each film was placed in 25 mL of 0.01 mM DPPH methanolic solution, incubated for 30 min at room temperature, and absorbance at 517 nm was measured. As a control, the same solution was measured but without any film samples. The DPPH radical scavenging activity was calculated using Equation (5).

Superoxide (O2 −) scavenging activity was assessed using the pyrogallol oxidation inhibition assay, following the methodology of Ye et al. with some modification [44]. Briefly, 100 mg of each film was incubated for 5 min in 3 mL of 50 mmol/L (pH 8.2) Tris-HCl buffer with gentle stirring. Then, 0.3 mL of 7 mM pyrogallol solution that was preheated to 25 ◦C was added and the mixture was allowed to react for exactly 4 min. To terminate the reaction 1 mL of 10 mM HCl was added and the absorbance was measured at 318 nm. The O2 − scavenging rate was calculated from the formula:

$$O\_2^- \text{ inhibition } (\%) = \left[ 1 - \frac{(A\_1 - A\_1')}{A\_0} \right] \times 100,\tag{6}$$

where *A*1 is the absorbance of the mixture in the presence of the sample, *A* 0 1 is the absorbance of water instead of the reaction agent, and *A*0 is the absorbance without the sample.

Hydroxyl (·OH) scavenging was assessed using the method of Ye et al. [44] with some modification. Film specimens (100 mg) were placed in a mixture of 1.5 mL of 5 µM ortophenantroline solution and 2 mL of phosphate buffer (pH 7.4, 0.05 M). Then, 1 mL of 7.5 mM FeSO4 solution was added, followed by 1 mL of 0.1% H2O2, and, finally, distilled water was added to bring the total volume to 10 mL. The reaction solution was incubated at 37 ◦C for 1 h, protected from light, and the absorbance was

measured at 536 nm. Ortophenatroline solution without H2O2 (replaced by 1 mL of methanol) served as a blank. The following formula was used to calculate hydroxyl scavenging:

$$\text{-OH inhibition (\%)}=\left[\frac{A\_2 - A\_1}{A\_0 - A\_1}\right] \times 100,\tag{7}$$

where *A*0 is the ortophenatroline solution without H2O2 addition, *A*1 is the absorbance without the sample, and *A*2 is the absorbance in the presence of the sample.

#### *2.11. Evaluation of Cytotoxicity*

In order to screen for potential cytotoxicity, extract tests and direct contact tests were carried out based on ISO 10993-5 using L929 murine fibroblasts [45]. Cells (passage 20–25) were maintained in complete growth medium: DMEM containing 10% FBS, 2 mM l-glutamine, 100 U/mL penicillin, and 100 µg/mL streptomycin. For each material, 8-mm discs were cut using a steel punch and sterilized using 20 min exposure to UV lamp in BSL-2 safety cabinet (Telstar Bio II Advance, Barcelona, Spain). Extracts were prepared by placing six 8-mm diameter discs of each material in a tissue culture plate and soaking in 1 mL of growth media (ratio: 3 cm2 /mL) for 24 h at 37 ◦C. Medical grade PCL (CAPA6340) was used as a negative control, nitrile glove (Mercator Nitrylex Classic, Kraków, Poland) served as a positive (toxic) control, and as a sham extract, 1 mL of media was added to an empty well. In parallel, 10,000 L929 cells were seeded per well of a 96-well plate and allowed to adhere and spread for 24 h. Next, the media was aspirated and replaced with 100 µL of extract, six technical replicates per material. After a further 24 h of culture, cell viability was assessed using an inverted light microscope (Delta Optical IB-100, Mi ´nsk Mazowiecki, Poland) and resazurin viability assay [46] using a fluorescent plate reader (Biotek Synergy HTX, Winooski, VT, USA) (excitation 540 nm, emission 590 nm).

For the direct contact assay, 30,000 L929 cells were seeded per well of a 48-well plate and allowed to adhere and spread for 24 h. Then, discs of each material (8 mm diameter) were placed directly on top of the cell monolayer (*n* = 5 discs per material). Cells were then maintained for another 24 h of culture and viability was assessed using an inverted light microscope and resazurin viability assay—without removal of discs—as described previously.

For both experiments, viability data obtained from resazurin assay was analyzed by subtracting blank signal (growth media only, no cells) from all other measurements and normalizing to sham-treated cells. Both the extract and direct contact assay were performed twice.

#### *2.12. Statistical Analyses*

Statistical comparisons were performed using Statistica version 10 (StatSoft Polska, Kraków, Poland). Differences between means were determined using analysis of variance (ANOVA) followed by Fisher's LSD post-hoc testing with a significance threshold of *p* < 0.05. All measurements were carried out in at least triplicate.

#### **3. Results and Discussion**

#### *3.1. Hydrodynamic Properties (Moisture Content, Water Solubility and Swelling Ratio)*

The MC, WS, and SR of the WPC/WPI films are summarized in Table 1. The MC of neat WPC film was 27.16 ± 1.31%, whereas the MC of neat WPI film was 24.31 ± 0.51%. Both increased significantly as melanin content increased. This observation is in line with the findings of other authors who used melanins to modify biopolymer-based films, such as chitosan and agar [14,19]. In contrast, Yang et al. [21] noticed that the MC of alginate/poly(vinyl alcohol) films decreased with increased melanin content, similarly as observed by Roy et al. for the case of cellulose/melanin films [22]. According to Roy et al., a significant increase in the MC of polymer/melanin films may be the result of reduced polymer network interactions and, as a consequence, greater accessibility of free hydroxyl groups to absorb more water molecules [14,19].

**Table 1.** Moisture content (MC), water solubility (WS) and swelling ratio (SR) of neat whey protein concentrate/isolate films (WPC/WPI) and melanin-modified films.

Values are means ± standard deviation. Means with different lowercase are significantly different at *p* < 0.05.

In terms of WS, melanin addition resulted in a significant decrease in WS for both types of films (*p* < 0.05). The WS of WPC-0.5% and WPI-0.5% was reduced by approximately 20.87% and 13.50%, respectively, as compared to the neat WPC and WPI films (*p* < 0.05). Previously, a WS reduction for gelatin films modified with fungal melanin was reported [38]. Finally, for the case of SR, we observed that the addition of melanin markedly increased the SR of films (*p* < 0.05). Further, for the case of WPC films, they had a much higher SR than WPI films: the SR values of WPC-0.5% and WPI-0.5% were 147.58 ± 18.04% and 469.47 ± 21.69%, respectively and are lower than those reported previously for agar/melanin films [19]. Interestingly, Roy et al. observed that SR of chitosan films was improved by the reinforcement of melanin nanoparticles [14], but in contrast, SR was reported to be decreased in the case of agar/melanin nanocomposite films [19]. In biopolymer-based films, SR primarily depends on the cross-linking, porosity, and nature of the materials [14,47]. Thus, in the present case, the cross-linking density of the melanin-modified films might be higher and, consequently, the porosity and SR of the samples may have increased [14].

#### *3.2. The Thickness, Mechanical and Thermal Properties*

The thickness, mechanical, and thermal properties of the samples are presented in Table 2. The modification with melanin did not affect the thickness of WPC or WPI films (*p* > 0.05), probably because of the small amount of melanin used. However, in studies by other authors who used melanin particles at higher concentrations, an increase of thickness was observed, due to higher dry mass content [19]. Here, the addition of melanin significantly enhanced the mechanical strength of WPC and WPI films (*p* < 0.05). The tensile strength (TS) of the WPI films was higher than the corresponding WPC films; the highest TS was observed for sample WPI-0.5% (6.13 ± 0.41 MPa). However, the TS of the WPC/WPI films was lower than that reported for agar/melanin [19], chitosan/melanin [14] and poly(lactic acid)/melanin films [18]. We also observed that the elongation at break (EB) of the modified films decreased in comparison to the control samples (*p* < 0.05). The increase in TS and decrease of EB can be attributed to strong hydrogen bonding (H-bonding) interactions between melanin and the polymer matrices, which improves the mechanical properties of the films. This observation is in good agreement with the results of other authors [14,19,22]. It has also been reported that reinforcement with a low percentage (0.025% and 0.05%) of fungal melanin improved the mechanical properties of poly(lactic acid)/melanin composite films; however, at higher melanin concentration (0.2%), the TS of the films decreased [18]. These results clearly indicate that melanins (synthetic or natural) can improve the mechanical properties of polymer-based films, although the effect of melanin is highly dependent on the type of polymer matrix used and melanin concentration, which play pivotal roles in the interand intramolecular interactions between the polymer chains in the film [14,18,19].

**Table 2.** Thickness, mechanical, and thermal characteristics of neat WPC/WPI and melanin-modified films.

Values are means ± standard deviation. Means with different lowercase are significantly different at *p* < 0.05.

In the present study, DCS measurements were also carried out to investigate the thermal characteristics of the films. Table 3 presents the melting temperatures (Tm) and melting enthalpies (∆Hm) of the samples. Compared to the neat WPC/WPI films, the addition of melanin markedly increased the Tm and ∆Hm of the modified films. These changes may be due to different polymer-water interactions as a result of modification with melanin [14]. For example, Dong et al. reported that the addition of synthetic and natural melanin to PVOH improved the thermal stability of the PVOH films [40]. However, there are also several reports that modification with melanin did not affect the thermal stability of agar [19], cellulose [22], and poly(lactic acid) films [18]. At the same time, in those reports, melanins were used as fillers. Thus, it can be reasonably concluded that the different effects of melanins on thermal stability may be due to different melanin sources, differences in the polymer matrix, as well as different methods of preparation of the films [13].

**Table 3.** Color parameters (L\*, a\*, b\*), total color difference (∆E), and yellowness index (YI) of neat WPC/WPI and melanin-modified films.

Values are means ± standard deviation. Means with different lowercase are significantly different at *p* < 0.05.

#### *3.3. Color*

The apparent color, the total color difference (∆E), and the yellowness index of the WPC/WPI films are presented in Table 3 and Figure 1. The neat WPI film was transparent and colorless, whereas neat WPC film showed a light yellowish color. Generally, WPC films were darker (lower L values) when compared with WPI films. The films with 0.1% and 0.5% melanin showed a yellowish-brown color and became darker as the concentration of melanin increased (*p* < 0.05). The redness and yellowness of the modified films also increased, due to the red-brown color of melanin. The yellowness index of WPC/WPI films increased significantly with increased melanin concentration (*p* < 0.05). As a result, the ∆E of the films was remarkably increased (*p* < 0.05) and ranged from 2.29 (WPC-0.1%) to 15.13 (WPC-0.5%). ∆E > 1 is considered perceptible to the human eye, so both concentrations of melanin in WPC/WPI films caused noticeable color changes [18]. Overall, the observed results are in good agreement with previous reports, where chitosan [14], agar [19], polypropylene/poly(butylene-co-terephthalate) [37], gelatin [38] and poly(lactic acid) [18] films were modified with various melanins.

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**Figure 1.** The visual appearance of neat and modified WPC/WPI films. **Figure 1.** The visual appearance of neat and modified WPC/WPI films. **Figure 1.** The visual appearance of neat and modified WPC/WPI films.

#### *3.4. UV-barrier Properties 3.4. UV-Barrier Properties 3.4. UV-barrier Properties*

UV-Vis light transmittance spectra of the neat and modified WPC/WPI films are shown in Figure 2. As can be seen, the neat WPC and WPI films were highly transparent to visible light at wavelengths greater than 380 nm and exhibited high transmittance for both UVA and UVB light. The transparency of WPI and WPC films decreased markedly by the addition of melanin and the decrease was concentration-dependent. These results clearly indicate that melanin, even at low concentration (0.1%) improves UV-light barrier properties of WPC, as well as WPI films. The decrease in the light transmittance of WPC and WPI films was mainly due to the absorption of UV light by melanin, similarly as in gelatin/melanin films [38]. However, when melanins were used as nanofillers, the UV barrier properties of materials were due to blocking of the light path, as was reported for poly(lactic acid)/melanin films [18] as well as for agar/melanin nanocomposite films [19]. In fact, in nature, melanins play a pivotal role in UV-protection, due to their strong UV-blocking properties resulting from the presence of phenolic and indole groups [22,37]. The high UV-barrier properties of melanin from watermelon seeds have also already been reported [10]. UV-Vis light transmittance spectra of the neat and modified WPC/WPI films are shown in Figure 2. As can be seen, the neat WPC and WPI films were highly transparent to visible light at wavelengths greater than 380 nm and exhibited high transmittance for both UVA and UVB light. The transparency of WPI and WPC films decreased markedly by the addition of melanin and the decrease was concentration-dependent. These results clearly indicate that melanin, even at low concentration (0.1%) improves UV-light barrier properties of WPC, as well as WPI films. The decrease in the light transmittance of WPC and WPI films was mainly due to the absorption of UV light by melanin, similarly as in gelatin/melanin films [38]. However, when melanins were used as nanofillers, the UV barrier properties of materials were due to blocking of the light path, as was reported for poly(lactic acid)/melanin films [18] as well as for agar/melanin nanocomposite films [19]. In fact, in nature, melanins play a pivotal role in UV-protection, due to their strong UV-blocking properties resulting from the presence of phenolic and indole groups [22,37]. The high UV-barrier properties of melanin from watermelon seeds have also already been reported [10]. UV-Vis light transmittance spectra of the neat and modified WPC/WPI films are shown in Figure 2. As can be seen, the neat WPC and WPI films were highly transparent to visible light at wavelengths greater than 380 nm and exhibited high transmittance for both UVA and UVB light. The transparency of WPI and WPC films decreased markedly by the addition of melanin and the decrease was concentration-dependent. These results clearly indicate that melanin, even at low concentration (0.1%) improves UV-light barrier properties of WPC, as well as WPI films. The decrease in the light transmittance of WPC and WPI films was mainly due to the absorption of UV light by melanin, similarly as in gelatin/melanin films [38]. However, when melanins were used as nanofillers, the UV barrier properties of materials were due to blocking of the light path, as was reported for poly(lactic acid)/melanin films [18] as well as for agar/melanin nanocomposite films [19]. In fact, in nature, melanins play a pivotal role in UV-protection, due to their strong UV-blocking properties resulting from the presence of phenolic and indole groups [22,37]. The high UV-barrier properties of melanin from watermelon seeds have also already been reported [10].

**Figure 2.** UV–Vis spectra of neat and modified WPC/WPI films. **Figure 2.** UV–Vis spectra of neat and modified WPC/WPI films.

#### *3.5. Antioxidant Activity*

The antioxidant activity of the films was determined by measuring the reducing power and radical (DPPH, ABTS, O2 −, OH) scavenging activity, as presented in Table 4. It was noticed that the antioxidant activity of the films was melanin concentration-dependent and increased significantly with increased melanin concentration (*p* < 0.05). The highest reducing power (1.186 ± 0.06) was measured for sample WPI-0.5%. The ABTS scavenging efficiencies of the neat WPI and WPI films were 72.02 ± 5.78% and 60.93 ± 4.50%, respectively, and an increase to 85.18 ± 1.74% and 96.87 ± 0.48%, respectively, was noticed when melanin concentration was 0.5%. A similar trend, but lower values, were observed in the case of DPPH and O2 − scavenging activity. On the other hand, the highest hydroxyl radical scavenging activity was observed for sample WPC-0.5% (39.08 ± 0.14%). In fact, the antioxidant activity of edible films is known to increase with the number of antioxidants [16,20,22,48]. Further, the antioxidant activity of the neat WPC/WPI films was expected, as they are reported to contain bioactive proteins, peptides, and amino acids with antioxidant capacity [28,31,49]. The antioxidant activity observed in the present report is also in good agreement with previous findings regarding agar [19], carrageenan [13], cellulose [22] and poly(lactic acid) [18] composite films modified with various melanins as fillers. Further, the antioxidant activity of melanin/gelatin films has been shown to increase when melanin is dissolved in a film-forming alkaline solution [38]. Likewise, it has already been observed that melanin from watermelon seeds has antioxidant activity [10]. In fact, it is well known that melanins act as effective antioxidants, because of intramolecular non-covalent electrons that can easily interact with free radicals and other reactive species [19]. Thus, the WPC/WPI melanin-modified films obtained here can be potentially used in active antioxidant packaging, to prevent oxidation of lipid-containing food or to preserve oxidation-sensitive food, as well to increase the shelf life of food [18,22,39]. Indeed, gelatin-based coatings modified with fungal melanin have been shown to have preservative activity against pork lard rancidity [39]. However, it is possible that the materials obtained here could also be used in biomedical applications, such as wound healing, due to their strong radical scavenging activity [31,50,51]. Wound healing is a complex process requiring an optimal balance between oxidative stress and antioxidant status. Typically, proper wound healing requires low levels of reactive oxygen species and low oxidative stress, while excessive oxidative stress can to impaired wound healing. As a result, antioxidants that can help control wound oxidative stress can improve wound healing [50,51].

**Table 4.** Reducing power (RP) and radical (DPPH, ABTS, O2 WPC/WPI and melanin-modified films.

−,

·OH) scavenging activity of neat

Values are means ± standard deviation. Means with different lowercase are significantly different at *p* < 0.05.

#### *3.6. FT-IR*

FT-IR is a commonly used technique to assess the miscibility and compatibility of biopolymers and additives, owing to its rapid and nondestructive nature [3,18,22]. Absorbance spectra of WPC-based and WPI-based films are presented in Figure 3. As can be seen the films have characteristic peaks at 3280 cm−1 attributed to C–H, N–H and/or O–H of WPC/WPI and melanin [10,27]. Characteristic CH3 and CH2 stretching peaks at approximately 2940 cm−1 and 2880 cm−1 were also observed. The peak centered at 1630 cm−1 can be attributed to Amide-I associated with C=O stretch and C–N vibrations; however, in the case of melanin-modified films, it also corresponds to the vibration of aromatic C=C of melanin [10]. Peaks in the region 1400–1550 cm−1 and 1200–1350 cm−1 can be assigned to Amide-II

*3.7. Water Contact Angle and WVTR*

polymer matrix.

(groups N-H) and to Amide III (with N–H bonds and C–N stretching), respectively [27]. The region from 750 to 1200 cm−1 was assigned to the absorption of glycerol (plasticizer), vibrations C–O and C–C bonds [27]. In the case of WPC and WPI films modified with melanin, slight variations in peak intensities and positions were observed, particularly at 3280 cm−1 , 2940 cm−1 , 2880 cm−1 and 1630 cm−1 . Overall, no substantial variations were noted in the functional groups of WPC/WPI modified films. The observed findings suggest that there were no structural changes in WPC and WPI films due to the addition of melanin. The small changes in intensities and minor shifts in the peaks are likely due to physical interactions (H-bonding, van der Waals force) between WPC/WPI and melanin, which is consistent with the findings of other authors [14,19,21]. (plasticizer), vibrations C–O and C–C bonds [27]. In the case of WPC and WPI films modified with melanin, slight variations in peak intensities and positions were observed, particularly at 3280 cm−1, 2940 cm−1, 2880 cm−1 and 1630 cm−1. Overall, no substantial variations were noted in the functional groups of WPC/WPI modified films. The observed findings suggest that there were no structural changes in WPC and WPI films due to the addition of melanin. The small changes in intensities and minor shifts in the peaks are likely due to physical interactions (H-bonding, van der Waals force) between WPC/WPI and melanin, which is consistent with the findings of other authors [14,19,21].

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respectively [27]. The region from 750 to 1200 cm−1 was assigned to the absorption of glycerol

**Figure 3.** FT-IR spectra of neat WPC/WPI and melanin-modified films. **Figure 3.** FT-IR spectra of neat WPC/WPI and melanin-modified films.

The results of water contact angle (WCA) measurements are presented in Table 5. The WCA of

neat WPC and WPI films was 27.67 ± 0.47° and 45 ± 0.00°, respectively. Generally, biopolymer films with a WCA of less than 65° are considered hydrophilic [14]. Concerning the effect of melanin addition, it was interestingly observed that the WCA of the films decreased significantly with

WCA of polymeric films due to their hydrophobic nature [14,18]. However, the difference might be attributed to the use of melanin particles in those studies, whereas in the present study melanin was used to modify the alkaline film-forming solution, which could result in other interactions in the

The water vapor permeability of the samples is also displayed in Table 5. It was observed that

in the case of WPC, as well as WPI, the WVTR decreased as a result of modification with melanin (*p* < 0.05). The observed results are in good agreement with previous reports, where various melanins

#### *3.7. Water Contact Angle and WVTR*

The results of water contact angle (WCA) measurements are presented in Table 5. The WCA of neat WPC and WPI films was 27.67 ± 0.47◦ and 45 ± 0.00◦ , respectively. Generally, biopolymer films with a WCA of less than 65◦ are considered hydrophilic [14]. Concerning the effect of melanin addition, it was interestingly observed that the WCA of the films decreased significantly with increasing melanin content to 14.33 ± 0.47◦ and 31 ± 0.00◦ for samples WPC-0.5% and WPI-0.5%, respectively. This observation is in contrast to the other reports that melanins generally increase the WCA of polymeric films due to their hydrophobic nature [14,18]. However, the difference might be attributed to the use of melanin particles in those studies, whereas in the present study melanin was used to modify the alkaline film-forming solution, which could result in other interactions in the polymer matrix.

**Table 5.** Water Contact Angle (WCA) and Water Vapor Transmission Ratio (WVTR) of neat WPC/WPI and melanin-modified films.

Values are means ± standard deviation. Means with different lowercase are significantly different at *p* < 0.05.

The water vapor permeability of the samples is also displayed in Table 5. It was observed that in the case of WPC, as well as WPI, the WVTR decreased as a result of modification with melanin (*p* < 0.05). The observed results are in good agreement with previous reports, where various melanins were used for the modification of alginate/poly(vinyl alcohol) [21], cellulose [22] and gelatin [38] films. However, there are also reports that the effect of melanin on WVTR depends on the concentration used. For instance, in the case of melanin added to poly(lactic acid) composite films, it was observed that WVTR decreased in the presence of low melanin content, but increased at high melanin content [18]. Likewise, in the case of agar/melanin nanoparticles composite films, the incorporation of low melanin-particle content did not result in WVTR changes, but at higher content, the WVTR increased. Whey protein-based materials are generally permeable or semipermeable to moisture penetration [25,52]. Although the WVTR of the melanin-modified films prepared here decreased (WVTR of WPC-0.5% and WPI-0.5%, 1483.53 ± 5.49 and 1490.49 ± 5.37 (g/(m2 × Day)) respectively), as compared to unmodified samples (WPC-C and WPI-C -1712.64 ± 7.46 and 1618.57 ± 6.23 (g/(m2 × Day)), respectively), they still exhibited weak water vapor barrier properties, lower than those reported in other studies [14,18,19,37].

#### *3.8. Cytotoxicity*

The cytotoxicity studies confirmed the non-toxic nature of the samples. As can be seen in Figure 4, after 24 h of exposure to extracts, robust cell growth was observed for all tested materials, similar to sham. Further, no marked differences in morphology were observed that could indicate a negative effect. The results of the resazurin cell viability assay, presented in Figure 5, were in good agreement with the microscopic observations. In all cases, cell viability was similar, near 90%—higher than the 70% threshold for cytotoxicity specified by the ISO 10993 norm.

*Materials* **2020**, *13*, x FOR PEER REVIEW 14 of 19

**Figure 4.** Representative micrographs of L929 fibroblasts. Panel (**A**): cells 24 h after seeding, but prior to the addition of extracts. Panel (**B**): Cells incubated for 24 h with the sham extract. Panels (**C**,**E**,**G**) present cells incubated with WPC extracts (WPC; WPC-0.1% and WPC-0.5%) for 24 h. Panels (**D**,**F**,**H**) present cells incubated with WPI extracts (WPI; WPI-0.1% and WPI-0.5%) for 24 h. The scale bar represents 200 μm. **Figure 4.** Representative micrographs of L929 fibroblasts. Panel (**A**): cells 24 h after seeding, but prior to the addition of extracts. Panel (**B**): Cells incubated for 24 h with the sham extract. Panels (**C**,**E**,**G**) present cells incubated with WPC extracts (WPC; WPC-0.1% and WPC-0.5%) for 24 h. Panels (**D**,**F**,**H**) present cells incubated with WPI extracts (WPI; WPI-0.1% and WPI-0.5%) for 24 h. The scale bar represents 200 µm. to the addition of extracts. Panel (**B**): Cells incubated for 24 h with the sham extract. Panels (**C**,**E**,**G**) present cells incubated with WPC extracts (WPC; WPC-0.1% and WPC-0.5%) for 24 h. Panels (**D**,**F**,**H**) present cells incubated with WPI extracts (WPI; WPI-0.1% and WPI-0.5%) for 24 h. The scale bar represents 200 μm.

**Figure 4.** Representative micrographs of L929 fibroblasts. Panel (**A**): cells 24 h after seeding, but prior

**Figure 5.** Viability of L929 fibroblasts after 24 h incubation with extracts. Data is normalized to sham extract. Dots represent technical replicates, grey bars represent median, and the red line indicates 70% **Figure 5.** Viability of L929 fibroblasts after 24 h incubation with extracts. Data is normalized to sham extract. Dots represent technical replicates, grey bars represent median, and the red line indicates 70% **Figure 5.** Viability of L929 fibroblasts after 24 h incubation with extracts. Data is normalized to sham extract. Dots represent technical replicates, grey bars represent median, and the red line indicates 70% viability, the threshold for cytotoxicity.

viability, the threshold for cytotoxicity. In terms of the direct contact assay, again robust cell growth was observed beneath the discs of In terms of the direct contact assay, again robust cell growth was observed beneath the discs of each material, visually similar to wells without discs (Figure 6). Likewise, no marked differences in In terms of the direct contact assay, again robust cell growth was observed beneath the discs of each material, visually similar to wells without discs (Figure 6). Likewise, no marked differences in

viability, the threshold for cytotoxicity.

each material, visually similar to wells without discs (Figure 6). Likewise, no marked differences in morphology were observed that could indicate a negative effect. Thus, the reactivity is graded as 0,

morphology were observed that could indicate a negative effect. Thus, the reactivity is graded as 0,

200 μm.

morphology were observed that could indicate a negative effect. Thus, the reactivity is graded as 0, according to ISO 10993. The resazurin viability assay indicated a modest reduction in viability for 4 of the 6 tested materials (Figure 7), which is likely due to mechanical damage to the monolayer caused by the physical presence of the disc on top of the cells. Meanwhile, the values above 100% may be due to an interaction between melanin and the resazurin reagent, as the assay was performed with the discs remaining in place to avoid the risk of further mechanical damage. To compare the viability data to the 70% viability threshold set by the ISO10993 standard, we fitted a linear model (estimated using ordinary least squares, using R (RStudio)) to predict (Normalized Viability (%)-Threshold (70%)) with Material. The model explains a significant and substantial proportion of variance (R2 = 0.96, F(6, 24) = 94.64, *p* < 0.01, adj. R2 = 0.95). For each material, the effect was positive, indicating viability values were above the threshold, and significant (*p* < 0.001). Both cell culture experiments were repeated and similar results were observed. Overall, the non-toxic nature of the films was expected, as no cytotoxic effects of whey proteins [29] nor melanins [14,53] have been reported. *Materials* **2020**, *13*, x FOR PEER REVIEW 15 of 19 according to ISO 10993. The resazurin viability assay indicated a modest reduction in viability for 4 of the 6 tested materials (Figure 7), which is likely due to mechanical damage to the monolayer caused by the physical presence of the disc on top of the cells. Meanwhile, the values above 100% may be due to an interaction between melanin and the resazurin reagent, as the assay was performed with the discs remaining in place to avoid the risk of further mechanical damage. To compare the viability data to the 70% viability threshold set by the ISO10993 standard, we fitted a linear model (estimated using ordinary least squares, using R (RStudio)) to predict (Normalized Viability (%)-Threshold (70%)) with Material. The model explains a significant and substantial proportion of variance (R2 = 0.96, F (6, 24) = 94.64, *p* < 0.01, adj. R2 = 0.95). For each material, the effect was positive, indicating viability values were above the threshold, and significant (*p* < 0.001). Both cell culture experiments were repeated and similar results were observed. Overall, the non-toxic nature of the films was expected, as no cytotoxic effects of whey proteins [29] nor melanins [14,53] have been reported.

**Figure 6.** Representative micrographs of L929 fibroblasts. Panel (**A**): Cells 24 h after seeding, but prior to the addition of discs. Panel (**B**): cells incubated for 24 h without disc. Panels (**C**,**E**,**G**) present cells beneath WPC discs (WPC; WPC-0.1% and WPC-0.5%) after 24 h of culture. Panels (**D**,**F**,**H**) present cells beneath WPI discs (WPI; WPI-0.1% and WPI-0.5%) after 24 h of culture. The scale bar represents **Figure 6.** Representative micrographs of L929 fibroblasts. Panel (**A**): Cells 24 h after seeding, but prior to the addition of discs. Panel (**B**): cells incubated for 24 h without disc. Panels (**C**,**E**,**G**) present cells beneath WPC discs (WPC; WPC-0.1% and WPC-0.5%) after 24 h of culture. Panels (**D**,**F**,**H**) present cells beneath WPI discs (WPI; WPI-0.1% and WPI-0.5%) after 24 h of culture. The scale bar represents 200 µm.

**Figure 7.** Viability of L929 fibroblasts after 24 h incubation in direct contact with discs. Data is normalized to wells without discs. Dots represent individual discs, grey bars represent median, and the red line indicates 70% viability, the threshold for cytotoxicity. **Figure 7.** Viability of L929 fibroblasts after 24 h incubation in direct contact with discs. Data is normalized to wells without discs. Dots represent individual discs, grey bars represent median, and the red line indicates 70% viability, the threshold for cytotoxicity.

#### **4. Conclusions**

**4. Conclusions**  This article explored the properties of WPC and WPI films modified with plant melanin derived from agro-industrial by-product: watermelon seeds. The properties of melanin-modified WPC/WPI films were compared to neat WPC and WPI films. The modification with melanin had a marked effect on the mechanical, antioxidant, hydrodynamic and barrier properties, but did not introduce cytotoxicity. We conclude that the obtained biofunctional WPC/WPI melanin-modified films with improved mechanical, UV-barrier, and water vapor barrier properties, along with strong antioxidant activity could be used for active food packaging or, potentially, biomedical applications, such as This article explored the properties of WPC and WPI films modified with plant melanin derived from agro-industrial by-product: watermelon seeds. The properties of melanin-modified WPC/WPI films were compared to neat WPC and WPI films. The modification with melanin had a marked effect on the mechanical, antioxidant, hydrodynamic and barrier properties, but did not introduce cytotoxicity. We conclude that the obtained biofunctional WPC/WPI melanin-modified films with improved mechanical, UV-barrier, and water vapor barrier properties, along with strong antioxidant activity could be used for active food packaging or, potentially, biomedical applications, such as wound dressings.

wound dressings. **Author Contributions:** Ł.Ł.: conceptualization, formal analysis, investigation, methodology, supervision, visualization, writing—original draft, and writing—review and editing; E.D., P.S.: visualization, investigation, methodology, formal analysis, and writing—review and editing; P.T., M.K.: investigation; M.Ś.: funding **Author Contributions:** Ł.Ł.: conceptualization, formal analysis, investigation, methodology, supervision, visualization, writing—original draft, and writing—review and editing; E.D., P.S.: visualization, investigation, methodology, formal analysis, and writing—review and editing; P.T., M.K.: investigation; M.S.: funding acquisition, ´ investigation; M.H.F.H.: methodology; A.B.: funding acquisition, formal analysis. All authors have read and agreed to the published version of the manuscript.

acquisition, investigation; M.H.F.H.: methodology; A.B.: funding acquisition, formal analysis. All authors have read and agreed to the published version of the manuscript. **Funding:** This research received no external funding.

ZUT) for access to the multi-functional plate reader.

doi:10.15376/biores.9.4.5774-5777.

**Funding:** This research received no external funding. **Acknowledgments:** The authors thank Karol Fijałkowski (Faculty of Biotechnology and Animal Husbandry, ZUT) for access to the multi-functional plate reader.

**Acknowledgments:** The authors thank Karol Fijałkowski (Faculty of Biotechnology and Animal Husbandry, **Conflicts of Interest:** The authors declare no conflict of interest.

#### **Conflicts of Interest:** The authors declare no conflict of interest. **References**

5. Maina, S.; Kachrimanidou, V.; Koutinas, A. A roadmap towards a circular and sustainable bioeconomy through waste valorization. *Curr. Opin. Green Sustain. Chem.* **2017**, *8*, 18–23, doi:10.1016/j.cogsc.2017.07.007.

© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

## *Article* **Alginate Biofunctional Films Modified with Melanin from Watermelon Seeds and Zinc Oxide/Silver Nanoparticles**

**Łukasz Łopusiewicz 1,\* , Szymon Macieja 1 , Mariusz Sliwi ´nski ´ 2 , Artur Bartkowiak 1 , Swarup Roy 3 and Peter Sobolewski 4**

**Abstract:** Bioactive films find more and more applications in various industries, including packaging and biomedicine. This work describes the preparation, characterization and physicochemical, antioxidant and antimicrobial properties of alginate films modified with melanin from watermelon (*Citrullus lanatus*) seeds at concentrations of 0.10%, 0.25% and 0.50% *w*/*w* and with silver and zinc oxide nanoparticles (10 mM film casting solutions for both metal nanoparticles). Melanin served as the active ingredient of the film and as a nanoparticle stabilizer. The additives affected the color, antioxidant (~90% ABTS and DPPH radicals scavenging for all melanin modified films) and antimicrobial activity (up to 4 mm grow inhibition zones of *E. coli* and *S. aureus* for both zinc oxide and silver nanoparticles), mechanical (silver nanoparticles addition effected two-fold higher tensile strength), thermal and barrier properties for water and UV-vis radiation. The addition of ZnONP resulted in improved UV barrier properties while maintaining good visible light transmittance, whereas AgNP resulted in almost complete UV barrier and reduced visible light transmittance of the obtained films. What is more, the obtained films did not have an adverse effect on cell viability in cytotoxicity screening. These films may have potential applications in food packaging or biomedical applications.

**Keywords:** melanin; watermelon; alginate; bioactive films; nanoparticles

#### **1. Introduction**

During the transport and storage of food products, it is important to ensure that the product is protected from any negative physical, chemical, or microbiological factors. In this way, packaging should delay food spoilage and slow down the loss of beneficial properties of the processed product, including both visual and nutritional qualities [1]. For example, wax coatings applied to the surface of fruits to extend their shelf life were the first materials used as a coating on food products [2]. In low-income countries, the main share of the total waste is those of organic origin, while in high-income countries, inorganic waste is the most common [3]. Potentially, some of these organic wastes (including agro-industrial by-products) can be used as a source of bioactive compounds such as phenolic compounds, dietary fiber, polysaccharides, vitamins, carotenoids, pigments and oils [4].

Plastics, due to their ease of processing and relatively low cost of raw materials, have become one of the most widely used materials in virtually all industries. Their annual production exceeds 300 million tons worldwide [5]. However, plastics present a problem with regard to waste management: much of plastic waste consists of packaging

**Citation:** Łopusiewicz, Ł.; Macieja, S.; Sliwi ´nski, M.; Bartkowiak, A.; Roy, S.; ´ Sobolewski, P. Alginate Biofunctional Films Modified with Melanin from Watermelon Seeds and Zinc Oxide/Silver Nanoparticles. *Materials* **2022**, *15*, 2381. https:// doi.org/10.3390/ma15072381

Academic Editor: Sandra Maria Fernandes Carvalho

Received: 17 February 2022 Accepted: 18 March 2022 Published: 23 March 2022

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2022 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

waste, of which up to 60% is food packaging. Furthermore, the majority of plastic waste (approximately 80%) ends up in landfills or pollutes land or water reservoirs, thus affecting entire ecosystems, from plants through to animals and humans [6–8]. Additionally, plastic production and disposal generate large amounts of greenhouse gases, which also has a significant negative impact on the environment. Therefore, finding less harmful alternatives to conventional plastics, such as renewable or biodegradable materials, is important to reduce the harmful impact of packaging on ecosystems [9].

One promising strategy involves biopolymers, polymers of natural origin: usually proteins or polysaccharides obtained from plants, bacteria, fungi, animals or seaweed. As biopolymers have extremely diverse properties, they can find applications in many different industries, including use in packaging, for example, of medical equipment, agricultural products, and other commodities [10,11].

Polysaccharides are a class of biopolymers that are particular abundant, including cellulose, chitin, xanthan gum, starch, carrageenan, and alginate [12]. Alginates are naturally occurring polymers consisting of linear copolymers of D-mannuronic and L-guluronic acids, linked by β-1,4-glycosidic bonds. Depending on the source, the two monomers can differ in proportion and be arranged in different, irregular patterns—these structural aspects affect the properties of alginate-based materials [13]. Example sources include seaweeds, mainly algae (*Laminaria hyperborea, L. digitata, L. japonica, Ascophyllum nodosum,* or *Macrocystis pyrifera*), whose alginic acid is a component of cell walls, or bacterial biosynthesis using *Azotobacter* and *Pseudomonas* bacteria [14]. Alginates have found applications in the textile, food, pharmaceutical, and chemical industries [15]. Importantly, the alginate backbone includes many free hydroxyl and carboxyl groups, making them highly amenable to modification and functionalization, using techniques such as oxidation, esterification and amidation. In this way it is possible to significantly influence the material properties, both physicochemical (such as solubility and affinity for water), as well as biological [16].

Melanins are a group of high molecular weight pigments, formed by the oxidative polymerization of indole and phenolic compounds [17]. In addition to imparting color to living organisms, melanins are also responsible for free radicals scavenging, immunostimulation, thermoregulation, protection from ultraviolet radiation, and have antiviral and antimicrobial properties [18–20]. Due to their antioxidant activity, melanins can be used to reduce metal compounds and thus synthesize metal nanoparticles. In contrast to traditional techniques for obtaining nanomaterials, the use of melanins can enable processes that are relatively fast and inexpensive, making them environmentally friendly [21,22]. Due to the aforementioned properties of melanins, they can carry out many active functions, which have resulted in numerous works describing the use of these pigments in combination with various polymers, including, although not limited to, bioactive food packaging [23].

Zinc oxide nanoparticles (ZnONP) are among the most commonly produced nanomaterials, next to titanium oxide and silicon dioxide NPs. As with other nanomaterials, the size and shape (and thus properties) of ZnONP depends on the preparation method. ZnONP, similar to titanium dioxide NP have a high ability to absorb ultraviolet radiation with a high band gap, making them transparent to visible light [24]. As a result, nano ZnO is commonly used in cosmetics, such as sunscreens, for protecting the skin against ultraviolet radiation. Additionally, ZnONP possess antimicrobial activity, making them suitable for the purification of water from microbial contaminants [25] or as antibacterial additives to paints and coatings [26,27]. Importantly, ZnONP are regarded as being safe (GRAS) by the United States Food and Drug Administration (USFDA, 21CFR182.8991). Combined, these properties make them very attractive for applications in active packaging systems [28].

Silver nanoparticles (AgNP) also possess potent antimicrobial properties, making them highly valued in the pharmaceutical and medical industries. As shown in numerous studies, AgNP are able to induce pore formation in bacterial cell membranes, most likely through the interaction of silver with sulfur present in membrane proteins. Further, when silver penetrates into the cell, it can interact with genetic material, causing its condensation inhibiting replication and gene expression [29–32]. Due to these properties, AgNP are used in various biomedical applications, such as in burn wound dressings and as catheter coatings. However, AgNP are also used in electronics, photonics, cosmetics, cleaning agents, and as disinfectants [26,33].

The antimicrobial and UV barrier properties of ZnONP and AgNP described above, along with potential other effects on the polymer matrix, provide prospective applications for these nanoparticles in active packaging materials. Nevertheless, the amount of nanoparticles loaded into the polymer matrix is crucial for the properties of the polymer, which as a consequence may deteriorate the quality of the packaging. As it has been shown for chitosan nanoparticles, the use of other than optimal amounts (smaller or larger) can affect the thermal, mechanical, barrier or optical properties [34].

In this work, our aim is to investigate the effect of the addition of melanin obtained from watermelon seeds, a food industry waste, on the properties of alginate films. The melanin was used as an additive directly or was used for synthesis of ZnONP and AgNP. To the best of our knowledge, there have been no reports on the modification of alginate films with either natural melanin or in combination with nanoparticles to improve the functionality of these materials. The chemical composition of the films after incorporation of melanin and nanoparticles was examined by FT-IR spectroscopy. In addition, the effect of melanin on the color, hydrodynamic and optical properties of the films was evaluated. Moreover, their potential bio(functionality) was assessed by evaluating their mechanical and barrier properties, as well as antioxidant and potential cytotoxicity in vitro.

#### **2. Materials and Methods**

#### *2.1. Materials and Reagents*

Calcium chloride, hydrogen peroxide, disodium phosphate, monosodium phosphate, 1,1 diphenyl-2-(2,4,6-trinitrophenyl) hydrazyl (DPPH), 2,2'-azino-bis(3-ethylbenzothiazoline-6-acid) sulfonic acid (ABTS), potassium persulfate, potassium hexacyanoferrate (III), trichloroacetic acid, iron (III) chloride, iron (II) sulfate, tris (hydroxymethyl)aminomethane, pyrogallol, sodium alginate (Mw = 1,450,000), and *o*-phenanthroline were purchased from Sigma-Aldrich (Darmstad, Germany). Glycerol, ammonia water, hydrochloric acid, sodium hydroxide, chloroform, ethyl acetate, ethanol and methanol were from Chempur (Piekary Sl ˛askie, ´ Poland). Zinc (II) nitrate, peptone water and MacConkey medium were from Scharlau Chemie (Barcelona, Spain). Silver (III) nitrate was from POCh (Gliwice, Poland). Chapman's medium was from Merck (Dramstadt, Germany). All reagents were of analytical grade.

The microorganisms used to evaluate the antimicrobial properties of the films were obtained from the American Type Culture Collection (ATCC). The strains used were *Escherichia coli* ATCC8739 and *Staphylococcus aureus* ATCC12600.

For in vitro cytotoxicity studies, murine fibroblast cell line (L929), as well as all cell culture reagents: Dulbecco's Modified Eagle Medium (DMEM), fetal bovine serum (FBS), L-glutamine, penicillin, streptomycin, and all other cell culture reagents were purchased from Sigma-Aldrich (Pozna´n, Poland). All sterile, single-use cell culture plasticware was purchased from VWR (Gda ´nsk, Poland).

#### *2.2. Preparation of Alginate Films*

Fresh Crimson Sweet watermelons (*Citrullus lanatus*) were purchased from a local market (Szczecin, Poland). Melanin isolation and purification was performed as previously described [35]. Briefly, watermelon seeds (5 g) washed in distilled water and dried were immersed in 50 mL of 1 M NaOH for 24 h with shaking (150 rpm, 50 ◦C). The resulting mixture was then centrifuged (6000× *g* rpm, 10 min) and the separated supernatant was brought to pH 2.0 by adding 1 M HCl and followed by centrifugation (6000× *g* rpm, 10 min). The resulting precipitate was hydrolyzed with 6 M HCl (90 ◦C, 2 h), centrifuged again (6000× *g* rpm, 10 min) and washed five times with distilled water. Finally, the obtained precipitate was washed with chloroform, ethyl acetate and ethanol, dried and ground in a mortar. To prepare melanin containing films, distilled water (400 mL) was poured into

500 mL bottles and melanin was added in appropriate amounts (0.008 g, 0.02 g and 0.04 g) in order to obtain melanin concentrations of 0.10%, 0.25% and 0.50% (*w*/*w*), respectively. Ammonia water (2 mL) was added to all samples to create an alkaline environment that facilitated melanin dissolution. The bottles were placed on a shaker overnight to completely dissolve the melanin. The resulting solutions were pressure filtered to separate insoluble residues. The solution bottles were then placed on magnetic stirrers and 8 g of alginate was slowly added to each bottle. They were then placed back on the shaker overnight at 60 ◦C to completely dissolve the alginate. Glycerol, a plasticizing agent, was added (30% (*w*/*w*) relative to the amount of alginate used) to the solutions and stirred on a magnetic stirrer for 10 min. A control sample without melanin was prepared in the same fashion. To prepare films, alginate solutions were poured into square polystyrene plates (120 mm × 120 mm) at 40 g of solution per plate and dried at 40 ◦C for 48 h. All assays were performed in eight replicates.

Samples of AgNP or ZnONP were prepared using the same general procedure, however, with the addition of silver nitrate or zinc nitrate in an aqueous solution of melanin. For this purpose, aqueous solutions of silver nitrate or zinc nitrate were slowly added dropwise to the melanin solutions using a pipette until 10 mM was obtained in the film-forming solution. The resulting solutions were then incubated at 90 ◦C for 1 h. The solutions were cooled to room temperature before adding alginate (8 g). To prepare the films, the alginate solutions were poured onto square polystyrene plates (120 mm × 120 mm) at 40 g of solution per plate and dried at 40 ◦C for 48 h. All experiments were performed in eight replicates.

#### *2.3. Characterization on Nanoparticles*

NP purification was performed prior to performing the analyses. For this purpose, the mixtures were centrifuged (ZnONP mixtures at 5000 rpm and AgNP at 14,000 rpm) for 5 min and then the resulting precipitate was washed with distilled water. The procedure was repeated three times.

Samples of mixtures of melanin with AgNP or ZnONP were filtered using syringe filters (pore size 0.45 µm) and 1 mL of the obtained filtrates analyzed using UV-Vis light absorption spectroscopy (Thermo Scientific (Waltham, MA, USA) Evolution 220 UV-Vis spectrophotometer). The spectra were collected over the wavelength range of 300–800 nm, with a resolution of 1 nm. Melanin-nanoparticles mixture were also dried overnight at 40 ◦C and the resulting powder was collected for chemical composition analysis using a Perkin Elmer Spectrum 100 FT-IR spectrophotometer (Waltham, MA, USA). The obtained powders were measured directly using attenuated total reflection (ATR). The spectra were recorded over a wavelength range of 650–4000 cm−1 , with a resolution of 1 cm−1 .

Dynamic light scattering (DLS) measurements were performed using Malvern Nanosizer ZS instrument (Worcestershire, UK) with a He-Ne laser source (633 nm). For each sample, three measurements were performed at each of two sample dilutions. For ZnONP, after purification, each pellet was re-dispersed in 2 mL of saline, yielding homogenous latté-colored suspensions. These suspensions were then further diluted 1:80 and 1:200 for measurements. For AgNP, after purification, pellets were dispersed in 1 mL of water, yielding pale dispersions, ranging from pink (0.5% MEL) to purple (0.1% MEL). These dispersions were measured neat, as well as after further 1:1 dilution with water.

Each DLS measurement was performed at 25 ◦C after 2 min of temperature equilibration and consisted of 10–15 runs, at 173◦ scattering angle, and using automatic attenuation to ensure count rates <500 kcps. Data presented consist of hydrodynamic diameter (intensityweighted average, "z-average" diameter) and polydispersity index (PDI) obtained from cumulant analysis using Malvern Zetasizer software v3.30.

#### *2.4. Biocomposite Film Characterisation*

#### 2.4.1. Determination of Moisture Content and Water Solubility

Moisture content (MC) was determined as the change in film (2 cm × 2 cm) weight after drying at 105 ◦C for 24 h. Water solubility (WS) was tested by adding pre-weighed film fragments (2 cm × 2 cm) to 30 mL of distilled water in conical tubes and stirring overnight. The samples were then centrifuged and the water was removed from the precipitate using a pipette. The samples were dried overnight at 60 ◦C and finally weighed again. Each material was tested in three replicates and mean values were calculated. The solubility of the films in water was calculated using the following formula:

$$S\ \left(\%\right) = \frac{W\_1 - W\_2}{W\_1} \times 100\tag{1}$$

where *W*1 is the initial and *W*2 the final weight of the films.

#### 2.4.2. Thickness, Mechanical, and Thermal properties of Alginate Films

Film thickness was measured using an electronic thickness gauge (Dial Thickness Gauge 7301, Mitoyuto Corporation, Kangagawa, Japan) with an accuracy of 0.001 mm. Each sample was measured 10 times at randomly selected points and the results were averaged.

The tensile strength and elongation at break of the films were assessed using a Zwick/Roell 2.5 Z static testing machine (Ulm, Germany). The tensile clamp spacing was 25 mm and the head travel speed was 100 mm/min.

Differential scanning calorimetry (DSC) was used to assess thermal properties (DSC 3, Mettler-Toledo LLC, Columbus, OH, USA) using sequential heat-cool-heat cycles over a temperature range of 30–300 ◦C at *φ* = 10 ◦/min, under nitrogen flow (50 mL/min).

#### 2.4.3. Water Vapor Transmission Rate (WVTR) of Films

The water vapor permeability test (WVTR) was carried out using a gravimetric method: moisture sorption by pre-dried calcium chloride (9 g) was examined in containers tightly covered with test films (8.9 cm2 ). The containers were weighed daily over a period of three days (starting at day zero every 24 h until day three of the start of the analysis) to monitor the weight gain of calcium chloride and thus the water vapor permeability of the films. Each material was tested in four replicates and mean values for each day were calculated to express WVTR in g/(m2 × day) [36].

#### 2.4.4. Spectral Analysis of Films

UV-Vis spectra of films were measured using a Thermo Scientific (Waltham, MA, USA) Evolution 220 UV-vis spectrophotometer. Strips of films, with a surface area matching the quartz cuvette (5.5 cm × 1 cm), were placed along with a cuvette in the apparatus and spectra were recorded over a wavelength range of 300–800 nm, with a resolution of 1 nm.

The chemical composition of the obtained films was also evaluated using a Perkin Elmer Spectrum 100 FT-IR spectrophotometer (Waltham, MA, USA). Pieces of the films were measured directly, in ATR mode (32 scans per sample), and spectra were recorded over a wavelength range of 650–4000 cm−1 , with a resolution of 1 cm−1 .

#### 2.4.5. Film Color Analysis

The effect of melanin and silver or zinc nanoparticles on the color of the obtained films was assessed using a colorimeter (CR-5, Konica Minolta, Tokyo, Japan). Each sample was tested 10 times, at randomly selected points. The results (mean ± standard deviation) were expressed as L\*, a\* and b\* parameters. In addition, ∆*E* (color difference) and *YI* (yellowing index) were calculated for comparison with unmodified alginate films, using the following equations:

$$
\Delta E = \left[ \left( L\_{\text{standard}} - L\_{\text{sample}} \right)^2 + \left( a\_{\text{standard}} - a\_{\text{sample}} \right)^2 + \left( b\_{\text{standard}} - b\_{\text{sample}} \right) \right]^{0.5} \tag{2}
$$

$$YI = 142.86b \cdot L^{-1} \tag{3}$$

#### 2.4.6. Antioxidant Potential of Films

The antioxidant activity was measured by reducing power and free radical scavenging activity (ABTS+ , DPPH, and superoxide (O − 2 ) radicals) techniques as previously described [37], with minor modifications. For each film, two samples were tested and each measurement was performed in triplicate.

To assess reducing power, 100 mg film samples cut into pieces were placed in 1.25 mL of phosphate buffer (0.2 M, pH 6.6) and then 1.25 mL of 1% potassium ferricyanide solution was added. The samples were incubated at 50 ◦C for 20 min, after which 1.25 mL of trichloroacetic acid was added. The samples were then centrifuged at 1107 RCF for 10 min. From each sample, 1.25 mL of supernatant was taken, diluted with an equal volume of distilled water, and 0.25 mL of 0.1% ferric chloride solution was added. Finally, the absorbance of the samples was measured at 700 nm.

Free radical scavenging activity assay was performed against ABTS+ , DPPH, and superoxide (O − 2 ) radicals. The ABTS+ radical was induced prior to the addition of the film by mixing 5 mL of 7 mM ABTS with 2.45 mL of potassium persulfate and allowing it to stand overnight at room temperature, in the dark. ABTS+ solutions were then prepared by diluting with ethanol to an absorbance of 0.7. For each test, 100 mg of film was added to 10 mL of ABTS+ solution and incubated for six minutes in the dark. As negative control, polypropylene film with the same size as the samples was used, whereas the prepared ABTS+ solution without any film was used as a blank control. The mixtures were incubated under identical conditions. The absorbance of the samples was measured at 734 nm and the free radical scavenging activity was calculated using the following equation:

$$\text{Free radical saving activity (\%)} = \frac{A\_{\text{sample}} - A\_{\text{control}}}{A\_{\text{sample}}} \times 100\tag{4}$$

where *Asample* is the absorbance of the ABTS+ solution with the addition of the tested films, and *Acontrol* is the absorbance of the blank ABTS+ solution.

For DPPH radical scavenging activity tests, 100 mg of each film was placed in 10 mL of 0.01 mM DPPH in methanol and incubated in the dark for 30 min. As negative control, polypropylene film with the same size as the sample was used, whereas the prepared DPPH solution without any film was used as a blank control. The mixtures were incubated under identical conditions. Absorbance was measured at 517 nm and free radical scavenging activity was calculated using the same equation as for ABTS.

Superoxide (O − 2 ) radical scavenging activity was evaluated using the pyrogallol oxidation inhibition assay. For this purpose, 100 mg of each film was incubated for 5 min in 3 mL of 50 mmol/L Tris-HCl buffer (pH 8.2), with gentle stirring every minute. Then, 0.3 mL of pyrogallol was added. After exactly four minutes, the reaction was stopped by adding 1 mL of 10 mM HCl and the absorbance was immediately measured at 318 nm. As negative control polypropylene film was used. The radical scavenging activity was calculated using the following equation:

$$\text{CO}\_2^- \text{ inhibition (\%)} = \left[1 - \frac{A\_1 - A\_1^{\prime}}{A\_0}\right] \times 100 \tag{5}$$

where *A*1 is the absorbance of the reaction mixture with the addition of the sample, *A*'1 is the absorbance of the reaction mixture with water instead of pyrogallol and *A*0 is the absorbance of the reaction mixture without addition of test samples.

#### 2.4.7. Antimicrobial Activity

The antimicrobial properties were investigated by measuring the inhibition of microbial growth on plate count agar (PCA) in the presence of discs made of test films. For *Escherichia coli*, MacConkey agar was used, while for *Staphylococcus aureus*, Chapman agar

was used. After a day of culture, single colonies were picked from the media and added to sterile peptone water prepared according to the manufacturer's instructions until an optical density of 0.5 McFarland was achieved. The microorganisms were then seeded onto PCA plates and two discs (2 cm in diameter) of each film were placed on top. After 24 h of culture, the zone of growth inhibition was measured.

#### 2.4.8. Evaluation of Cytotoxicity

Cytotoxicity screening was performed using a direct contact assay, as described previously [37]. L929 cells were maintained in culture using DMEM containing 10% FBS, 2 mM l-glutamine, 100 U/mL penicillin, and 100 µg/mL streptomycin. For each test film, discs (8 mm in diameter) were cut using a steel punch, followed by sterilization with 20-min UV exposure in a BSL-2 safety cabinet (Telstar Bio II Advance, Barcelona, Spain).

For each experiment, L929 cells (passage 7–16) were trypsinized, counted using a hemocytometer, and seeded in the wells of 48-well plates (30,000 cells per well). After 24 h of incubation to permit the cells to adhere and spread, the media was aspirated and discs of each test film (5–6 samples each) were placed directly onto the cell monolayer—one well at a time. Manipulation of the thin discs was performed using a sterile 21 G needle: discs were "speared" vertically (from above), lifted, positioned in place in the well, and released by a gentle twist of the needle. Afterwards, 0.25 mL of fresh media was gently pipetted on top. As a sham control for normalization, wells with cells were aspirated, although no disc was placed, prior to adding 0.25 mL of fresh media. After a further 24 h of culture, cell viability was assessed by inverted light microscopy (Delta Optical IB-100, Minsk Mazowiecki, Poland) and using the resazurin assay [38]. Note that it was not possible to remove the discs, as they partially dissolved and in the case of samples containing nanoparticles, the nanoparticles interfered with the resazurin assay. Fluorescence plate reader measurements (Biotek Synergy HTX, Winooski, VT, USA, excitation 540 nm, emission 590 nm) were converted to normalized cell viability as percentage of sham by first subtracting the mean signal of six blank wells (no cells) from all values, followed by dividing by the mean signal of the sham wells (n = 6).

#### *2.5. Statistical Analyses*

All analyses were made at least in triplicate. Statistical analysis was performed using Statistica version 13 software (StatSoft Poland, Krakow, Poland). Differences between means were determined by analysis of variance (ANOVA) followed by Fisher's post hoc LSD test at a significance threshold of *p* < 0.05.

#### **3. Results and Discussion**

#### *3.1. Characterization of Nanoparticles*

In order to investigate the quality of obtained nanoparticles, UV-Vis analysis was carried out. UV-Vis spectra of ZnONP are shown in Figure 1A. The absorption maximum was at 307 nm. This value is shifted towards the UV-B radiation, as compared to the results obtained by Roy and Rhim [22], who noted a maximum at 365 nm for ZnO nanoparticles prepared using melanin.

UV-Vis spectra of AgNP are shown in Figure 1B. The maximum absorption peak can be observed at 420, 421, and 361 nm for 0.10% MEL + nAg, 0.25% MEL 0.25% + nAg, and 0.50% MEL + nAg, respectively. These results are similar to data obtained by Shankar and Rhim [39], as well as Vilmala et al. [40].

In order to investigate the presence of characteristic bonds in the samples FTIR analysis was performed. Both silver and ZnONP samples may contain melanin used in the synthesis of these nanoparticles, which was assessed using FTIR. Figure 2 shows the FTIR spectra of the AgNP obtained from the 0.50% MEL + nAg sample. Several distinct peaks can be seen. The stretching vibration of –OH bonds (phenols) at 3200 cm−1 are likely from the melanin present in the sample, along with the peaks at 1275 cm−1 and 1033 cm−1 corresponding to

C–O stretching, and 800 cm−1 , which can be attributed to aromatic groups. These results are similar to measurements of AgNP biosynthesized using plant extracts [41]. obtained by Roy and Rhim [22], who noted a maximum at 365 nm for ZnO nanoparticles prepared using melanin.

was at 307 nm. This value is shifted towards the UV-B radiation, as compared to the results

*Materials* **2022**, *15*, 2381 8 of 25

**Figure 1.** UV-Vis spectra of obtained zinc oxide nanoparticles (**A**), silver nanoparticles (**B**) samples, UV-Vis spectra of *C. lanatus* melanin at different concentrations for comparison (**C**).

**Figure 1.** UV-Vis spectra of obtained zinc oxide nanoparticles (**A**), silver nanoparticles (**B**) samples,

UV-Vis spectra of AgNP are shown in Figure 1B. The maximum absorption peak can be observed at 420, 421, and 361 nm for 0.10% MEL + nAg, 0.25% MEL 0.25% + nAg, and 0.50% MEL + nAg, respectively. These results are similar to data obtained by Shankar and

In order to investigate the presence of characteristic bonds in the samples FTIR analysis was performed. Both silver and ZnONP samples may contain melanin used in the synthesis of these nanoparticles, which was assessed using FTIR. Figure 2 shows the FTIR spectra of the AgNP obtained from the 0.50% MEL + nAg sample. Several distinct peaks can be seen. The stretching vibration of –OH bonds (phenols) at 3200 cm−1 are likely from the melanin present in the sample, along with the peaks at 1275 cm−1 and 1033 cm−1 corresponding to C–O stretching, and 800 cm−1, which can be attributed to aromatic groups. These results are similar to measurements of AgNP biosynthesized using plant extracts

UV-Vis spectra of *C. lanatus* melanin at different concentrations for comparison (**C**).

Rhim [39], as well as Vilmala et al. [40].

[41].

Figure 3 shows the FTIR spectra of ZnONP. Considerable differences in the intensity and position of the FTIR peaks can be noted for the individual samples. This is likely due to the different amounts of residual melanin due to the different concentrations used during preparation. The primary peaks occur in the spectral regions: 3500–3250 cm−1, 1600– 1550 cm−1, 1400–1300 cm−1, 1100–950 cm−1, and 900–750 cm−1. These results are in good agreement with data obtained by Roy and Rhim [22], who also used melanin for ZnONP synthesis. Figure 3 shows the FTIR spectra of ZnONP. Considerable differences in the intensity and position of the FTIR peaks can be noted for the individual samples. This is likely due to the different amounts of residual melanin due to the different concentrations used during preparation. The primary peaks occur in the spectral regions: 3500–3250 cm−1 , 1600–1550 cm−1 , 1400–1300 cm−1 , 1100–950 cm−1 , and 900–750 cm−1 . These results are in good agreement with data obtained by Roy and Rhim [22], who also used melanin for ZnONP synthesis.

In order to investigate average particle size, dynamic light scattering was used. Dynamic light scattering results of the obtained particles are presented in Table 1. For all ZnONP samples, unimodal, yet broad distributions were obtained, with a trend towards smaller particles with higher MEL content. At 0.50% MEL, the particles were the smallest, sub-1-micron in diameter and with the narrowest distribution (PDI = 0.266 ± 0.061). For the case of AgNP, particle sizes were markedly smaller, all <300 nm, and the effect of MEL concentration was different: the highest MEL concentration (0.50%) yielded the largest particles, although again with the narrowest distribution (PDI = 0.288 ± 0.037). Additionally, for the 0.10% MEL AgNP, a minor shoulder was observed (~10% of area) at ~60 nm diameter.

**Table 1.** Dynamic light scattering results: hydrodynamic diameter (z-avg) and polydispersity index (PDI). Data are expressed as mean ± standard deviation (SD) of six measurements, 10–15 runs each.

Values are means ± standard deviation. Means with different letters (a–c) are significantly different at *p* < 0.05 (compared to other variants within groups).

**Figure 3.** FTIR spectra of obtained zinc oxide nanoparticles samples. **Figure 3.** FTIR spectra of obtained zinc oxide nanoparticles samples.

#### In order to investigate average particle size, dynamic light scattering was used. Dynamic light scattering results of the obtained particles are presented in Table 1. For all *3.2. Hydrodynamic Properties (Moisture Content and Water Solubility) and Water Vapor Transmission Rate (WVTR)*

ZnONP samples, unimodal, yet broad distributions were obtained, with a trend towards smaller particles with higher MEL content. At 0.50% MEL, the particles were the smallest, sub-1-micron in diameter and with the narrowest distribution (PDI = 0.266 ± 0.061). For the case of AgNP, particle sizes were markedly smaller, all <300 nm, and the effect of MEL concentration was different: the highest MEL concentration (0.50%) yielded the largest particles, although again with the narrowest distribution (PDI = 0.288 ± 0.037). Additionally, for the 0.10% MEL AgNP, a minor shoulder was observed (~10% of area) at ~60 nm diameter. **Table 1.** Dynamic light scattering results: hydrodynamic diameter (z-avg) and polydispersity index (PDI). Data are expressed as mean ± standard deviation (SD) of six measurements, 10–15 runs each. **Sample Hydrodynamic Diameter (nm) PDI**  0.10% MEL + nZnO 1523 ± 201 a 0.476 ± 0.172 a In order to investigate the influence on hydrodynamic properties of the addition of melanin, ZnONP and AgNP, moisture content, water solubility and water vapor transmission rate have been examined. The moisture content and water solubility of the unmodified and modified alginate films are shown in Table 2. The moisture content of the unmodified alginate film was 11.61 ± 0.22%. The addition of melanin caused a significant increase to 14.93 ± 3.14% for ALG + 0.10% MEL film (*p* < 0.05). These values are similar to those reported for agar films with melanin [42]. While the effect of melanin addition on the moisture content of the films was consistent with the results of other researchers [22,37,43,44], there are also reports of the opposite effect of melanin [45,46]. These differences can be attributed to the different properties of the biopolymers used and the resulting different interactions between their functional groups with melanin, leading to an increase or decrease in the overall availability of free hydroxyl groups able to bind water molecules [46].

0.50% MEL + nZnO 988 ± 42 b 0.266 ± 0.061 a 0.10% MEL + nAg 230 ± 6 c 0.412 ± 0.023 a **Table 2.** Moisture content (MC), water solubility (WS) and Water Vapor Transmission Ratio (WVTR) of unmodified alginate films and films modified with melanin and melanin with nanoparticles.

0.25% MEL + nZnO 1167 ± 74 b 0.312 ± 0.104 a

Values are means ± standard deviation. Means with different lowercase (a–c) are significantly different at *p* < 0.05 (compared to control), means with different uppercase (A–B) are significantly different at *p* < 0.05 (compared to other variants within groups).

In contrast to melanin, the addition of AgNP caused a decrease in MC, to a range from 10.96 ± 1.42 to 11.61 ± 0.29%, which is comparable to the results of Rhim et al. [47]. However, the addition of ZnONP to the films did not affect the MC of the obtained films. Interestingly, Kotharangannagari and Krishnan [48] observed a decrease in moisture content with increasing amount of zinc oxide added to films made of starch with lysine and various concentrations of ZnONP. This again indicates that the specific interactions between the biopolymer and the nanoparticle drive film affinity for moisture.

The water vapor barrier properties of the tested films are presented in Table 2. Compared to control alginate films, the WVTR of all of the modified alginate films had lower WVTR, with the effect being more pronounced with increasing concentration in the melanin used. For MEL addition alone, the WVTR drops from 1589.70 ± 54.96 g/(m2 × Day) for neat alginate films to 1117.23 ± 285.85 g/(m2 × Day) for the highest melanin concentration. As reported by Bang et al. [49], the effects of melanin on water vapor barrier properties can be attributed to the interactions of melanin with the polymer matrix, especially with free hydrophilic chains. As a result, previous studies have found both positive effects of melanin on WVTR [49], as well as negative effects [36,37] and a variable effect dependent on the melanin concentration [44].

Compared to the addition of melanin, the presence ZnONP and AgNP resulted in a smaller change in WVTR through the films. At the highest concentration of melanin, the WVTR was 1463.20 ± 116.17 and 1229.34 ± 57.05 g/(m2 × Day), for ZnONP and AgNP, respectively. However, both of these values remain lower than control alginate. For the case of ZnONP, the reduced effect may be attributed to their much larger size (~1 µm). In previous work, Roy and Rhim [22] studied carrageenan films with the addition of ZnONP synthesized with melanin, similar to this work. They observed that the addition of ZnONP nanoparticles increased the barrier properties of the film. Likewise, Kanmani and Rhim [50] as well as Shankar et al. [51] obtained similar results. However, for PLA, Chu et al. [52] observed an increase in water permeability through films after adding ZnONP, although no significant effect of the addition of AgNP. Further, Shankar and Rhim [39] noted a slight increase in water vapor permeability after adding AgNP to agar films, whereas Rhim et al. [53], in a similar test system, noted a decrease in this parameter with increasing silver nanoparticle concentration. Overall, the water vapor barrier properties are highly dependent on the properties of the materials involved, their interactions, and the preparation, and dispersion of the nano-additives used.

In terms of water solubility, the addition of melanin reduced solubility from 79.70 ± 11.26% to 73.54 ± 14.05% for ALG + 0.10% MEL and 75.68 ± 9.64% for ALG + 0.50% MEL, although the differences were not significant (*p* > 0.05). A similar effect of melanin was previously described for whey protein isolate and concentrate films, as well as for gelatin films [33,47]. However, Roy et al. [43] noted an inverse relationship for chitosan films, attributed to interactions of melanin with polymer chains.

Meanwhile, films with AgNP had lower solubility (61.92 ± 1.68% for ALG + 0.10% MEL + nAg) than the controls, however, as the amount of melanin in the film increased, the solubility significantly increased, to 69.79 ± 2.19% (*p* < 0.05). The same relationship was also observed for the samples with ZnONP: a trend of increasing water solubility from 69.37 ± 3.86% to 73.23 ± 2.50% with increasing melanin concentration. Previously, Rhim et al. [45] reported an increase in the solubility of chitosan films after the addition of nanosilver, although the differences were not statistically significant.

#### *3.3. The Thickness, Mechanical and Thermal Properties*

The thickness, mechanical and thermal properties of the tested films are summarized in Table 3. In general, the addition of melanin did not have a significant effect on the thickness of the films. This is in good agreement with prior work with films made of whey protein concentrate/isolate [37], although not with those made of agar [42], gelatin [54], carrageenan [22] or polybutylene adipate terephthalate (PBAT) [49] where melanin addition

tended to increase film thickness. Meanwhile, for carboxymethyl cellulose (CMC) films, melanin had no effect on the film thickness [36].

**Table 3.** Thickness, mechanical, and thermal characteristics of unmodified and modified alginate films.

Values are means ± standard deviation. Means with different lowercase (a–c) are significantly different at *p* < 0.05 (compared to control), means with different uppercase (A–C) are significantly different at *p* < 0.05 (compared to other variants within groups).

Regarding mechanical properties of the films, the tensile strength (TS) increased for films with increasing melanin content from 5.97 ± 1.12 MPa for films with 0.10% (*w*/*w*) melanin content to 17.59 ± 3.91 MPa for films with the highest content. An upward trend was also be observed for the films with the addition of nanoparticles, although the effect was not as pronounced. The results are broadly similar, however lower in magnitude than those obtained by other authors [22,36,42,44,45,49,54,55]. Elongation at break (EB) also increased with increasing melanin content. However, previous studies suggest that the value of the elongation at break may increase with increasing melanin concentration up to a point, beyond which it begins to decline; the same relationship may also exist for the tensile strength [43,49,55].

Surprisingly, the addition of ZnONP decreased the values of both tensile strength (TS) and elongation at break (EB) to 6.10 ± 3.06% and 1.60 ± 0.32%, respectively. Previously, Roy and Rhim [22] had found no significant changes for carrageenan films after the addition of these nanoparticles. For non-biopolymers the literature is mixed: Mania et al. [56] reported an increase in both of these parameters for polyethylene films due to the action of nano ZnO, whereas Chu et al. [52] recorded a decrease in TS and an increase in EB for poly (lactic acid) (PLA) films.

In contrast to ZnONP, AgNP caused an increase in tensile strength to approximately 15 MPa and a decrease in elongation at break to about 2% for all samples. In contrast, Rhim et al. [53], did not observe significant changes in EB after adding these nanoparticles to agar films, however, TS values were lower at low concentrations of nanoparticles used and then increased with the increase in the amount of AgNP. However, Shankar and Rhim [39] showed a decrease in TS for agar films, with a simultaneous increase in EB value. Again, the literature is mixed for non-biopolymer PLA: Chu et al. [52] observed that TS decreased slightly and EB slightly increased, while Fortunati et al. [57] found that both parameters tended to decrease with AgNP addition.

Overall, the results obtained for alginate films and the results reported in the literature for various polymer matrices indicate that the addition of both melanin and silver and ZnONP can have different effects, i.e., positive, negative or no change in the described mechanical parameters, depending on the type of polymer used, the amount of melanin/nanoparticles used, as well as the methods and conditions of nanocomposites preparation.

Table 3 presents the melting temperatures (Tm) and melting enthalpies (∆Hm) of the films as measured by DSC. Compared to the neat alginate films, the addition of melanin decreased the Tm for films containing the highest concentration of melanin (from 127.5 ± 1.9 ◦C to 99.8 ± 1.5 ◦C), with ∆Hm also decreasing with increasing melanin concentration (from 106.0 ± 13.6 J/g to 66.7 ± 8.5 J/g). For both ZnONP and AgNP, Tm decreased, while ∆Hm increased, although the values of these parameters remained similar for individual trials. This is in contrast to previous work with WPC/WPI films [37], where modification with melanin increased their thermal stability. However, addition of melanin to agar [42], cellulose [55] or PLA [44] films did not affect these parameters.

#### *3.4. UV Barrier Properties*