Active Biodegradable Packaging Films Based on the Revalorization of Food-Grade Olive Oil Mill By-Products

Abstract

Synthetic packaging is being replaced by biodegradable packaging through the revalorization of food industry by-products. The olive oil (OO) industry, known for producing large quantities of antioxidant-rich by-products, can be a major supplier for sustainable packaging materials. This study aims to valorize a food-grade by-product (defatted flour, DF) from OO extraction produced using a zero-waste strategy that combines expeller press technology and supercritical CO₂ extraction. DF and its aqueous extract (DFE) were combined with carboxymethylcellulose (CMC) to create biodegradable bioactive packaging films. DF contains a high content of insoluble dietary fiber (28.4%) and total phenolic compounds (35,000 ppm), including oleuropein, elenolic acid, hydroxytyrosol, and tyrosol (4324, 3603, 1525, and 157 ppm, respectively). This study examined the effects of DF and DFE on the physicochemical and barrier properties of the films, as well as their capacity to delay oxidation in polyunsaturated fatty acid-rich oil. Films with DF and DFE contained high phenolic content (1500 and 1200 ppm, respectively), and their inclusion improved ultraviolet visible barrier capacity. Additionally, oil oxidation was slower when protected by DF- and DFE-based films than when protected with CMC film alone. This allows their use as protective packaging and potential carriers of bioactive oils to enhance the nutritional and functional qualities of packaged foods.

1. Introduction

The by-products generated in the production of olive oil (OO) represent a great challenge for the food industry; though they contain significant amounts of compounds with important biological activities, at the same time, they are polluting materials for the environment if its revaluation is not considered [1]. From the by-products generated annually in oil mills around the world, an enormous amount of phenolic compounds (PCs), such as oleuropein, hydroxytyrosol, and tyrosol, which have antioxidant, anti-inflammatory, antibacterial, and antiviral functions, are wasted, which may be beneficial for human consumption but whose consequences for the environment are devastating due to high concentrations of PCs that can decolorize rivers and inhibit water oxygenation and plant seed germination [2,3].

For these reasons, alternative OO extraction processes are necessary to eliminate water use, minimize environmental impact, increase the total amount of PC in the oil, and preserve the food quality of the pomace for other applications in the food industry. One way to achieve this is by the sequential extraction of olive oil (OO) from dehydrated olives, combining expeller press extraction in which obtained oil is of similar quality to that obtained by conventional methods [4,5], and supercritical CO₂ is used to remove oil from the press cake, yielding a completely defatted pomace or flour [6]. The oil extracted from the press cake has proven to have quality indexes within those established by legislation and a much higher PC content than conventional extra virgin olive oil (EVOO) [7]. The result of these processes is a dry pomace with a high PC content, whose food quality remains intact because it has not been in contact with organic solvents. Therefore, the possible application of this flour is its incorporation in food to increase its nutritional value or by combining it with biodegradable packaging to produce bioactive packaging. Recent studies have shown that films made from agricultural waste extracts, such as peanut skin, pink pepper, coffee, wine, or OO industries, plays an important part in active packaging technology, resulting in novel methods for extending shelf life and maintaining or monitoring the quality of foods [8,9].

Food packaging has undergone significant changes in recent years due to a number of reasons, not only to preserve the environment, but also to save energy, reduce manufacturing costs, extend the shelf life of foods, and protect their sensory and nutritional quality, as well as the health of consumers [10]. However, concerns have been raised about the potential adverse health effects of certain plastics used in packaging since the ingestion of harmful chemicals leached from them can disrupt endocrine function, while their accumulation in adipose tissue poses long-term health risks to humans [11].

Potentially, a biodegradable packaging can be made from polylactic acid (PLA), polyhydroxyalkonates (PHAs), polyesters, and starch blends [12]. Recently, interest in cellulose-derived materials has increased. Carboxymethyl cellulose (CMC), a water-soluble anionic polysaccharide, has emerged as a potential material for developing active packaging due to its film-forming properties, such as mechanical strength, barrier properties, transparency, and thermal stability [13,14]. The addition of by-products or aqueous plant extracts to CMC films can reduce gas permeability and also protect against light [15], which initiates lipid oxidation in various foods, especially those containing large amounts of polyunsaturated fatty acids (PUFAs).

In the present work, the elaboration of bioactive and environmentally friendly packaging biofilms based on carboxymethylcellulose by adding as functional ingredients the by-products obtained from the production of olive oil, such as the defatted flour and its aqueous extract, was carried out. The physicochemical characterization of these biofilms and their ability to protect against oxidation and to extend the shelf life of a polyunsaturated microalgae oil highly susceptible to oxidation were evaluated.

In addition, this highly valuable biological and technological by-product was obtained from a sequential olive oil extraction process using clean technologies (expeller and supercritical CO₂) that promote the sustainability of the extraction processes and can be considered a “zero waste” process since the elimination of the use of water, reducing liquid waste and maximizing the utilization of the solid by-product, occurs.

2. Materials and Methods

2.1. Materials

Cornicabra olives harvested at the end of the 2020/2021 growing season were supplied by the Arganda oil cooperative, located in Arganda del Rey, Madrid province, Spain. The defatted flour (DF) was obtained from the sequential extraction of olive oil (OO) carried out following a methodology previously developed by our research group, which consists of a previous dehydration and destoning of the olive and the subsequent extraction of the oil by combining an expeller press to obtain a first OO and a press cake that is then used to extract a second OO by supercritical fluid (SCF) technology with carbon dioxide (CO₂) with a purity of 99.9% (Carburos Metalicos; Barcelona, Spain) and thus finally obtain a totally defatted flour [16]. Microalgae oil sourced from the marine microalgae Schizochytrium sp. was supplied by Progress Biotech (Capelle aan den Ijssel, The Netherlands). Solvents used were methanol (MeOH) and chloroform (CLF), which were supplied by Macron (Avantor Performance Material, Center Valley, PA, USA). Reagents used for determinations such as the Folin–Ciocalteu reagent and sodium carbonate (Panreac, Barcelona, Spain) and hydroxytyrosol (HT) standard ≥ 98% were supplied by Seprox Biotech (Madrid, Spain). Food-quality glycerol and pure standards of oleic squalene ≥ 98%, α-tocopherol ≥ 98%, caffeic acid ≥ 99%, syringic acid ≥ 99%, apigenin ≥ 95%, and oleuropein aglycone≥ 99% were supplied by Sigma-Aldrich (St. Louis, MO, USA), and sterol ≥ 99% was supplied by 137 Vita-Solar Biotechnology Co., Ltd. (Xi’an, China). Peroxide reagents were supplied by CDR FoodLabFat (Florence, Italy).

2.2. Olive Defatted Flour Characterization

2.2.1. Humidity and Particle Size

The total moisture content of defatted flour (DF) was measured using a 0.1 mg precision analytical balance by determining the difference in weight after drying 10 g of flour in an oven at 105 °C to constant weight. For the particle size determination, 50 g of DF was introduced into an electric sieve shaker with four sieves of different pore sizes (>500 μm, 500–250 μm, 250–100 μm, 100–45 μm, and <45 μm) along with a container to collect particles smaller than 45 μm. Samples were processed in the sieve shaker with ultrasonic vibration for 6 min at an amplitude of 1.8 mm with cycles of 9.0 s on and 1.0 s off. Each fraction was then collected and weighed. All measurements were performed in duplicate.

2.2.2. Total Fat Content

The total fat content (TFC) of DF was determined according to the Folch method [17]. In total, 200 mL of CLF:MeOH (2:1 v/v) was added to 10 g of DF and homogenized using an Ultraturrax T18 basic IKA (Staufen, Germany) at 12,500 rpm for 2 min. The homogeneous mixture was then magnetically stirred at 900 rpm for 60 min. The organic phase was mixed with distilled water in a 1:5 ratio, vortexed for 1 min, and centrifuged at 10,000 rpm for 5 min after vacuum filtration. The lower phases were collected and the solvent was evaporated in a rotary evaporator (Büchi B-480, Uster, Switzerland) at 40 °C, under 10 mbar until constant weight.

2.2.3. Lipid Characterization by Gas Chromatography

Gas chromatography (GC) analyses of the total fat extract obtained by the Folch method of DF were carried out in a gas chromatograph, Agilent 7820A (Agilent Technologies, Santa Clara, CA, USA), with on-column injection coupled to a flame-ionization detector (FID) according to Torres et al. [18]. An HP-5MS capillary column was used, 5% phenyl methyl silicone (length: 7 m, internal diameter: 0.25 mm, and thickness: 0.25 µm). The injection volume was 0.1 µL, with injector and detector temperatures at 50 °C and 340 °C, respectively. The temperature program started at 60 °C, and increased to 250 °C at 42 °C min⁻¹ (held for 20 min), and then to 340 °C at 25 °C min⁻¹ (held for 35 min). The lipid classes, fatty acids (FAs), triglycerides (TAGs), diglycerides (DAGs) and monoglycerides (MAGs), and minor compounds (squalene, α-tocopherol, sterols, and sterol esters) were identified by comparing their retention times with those of reference standards. Quantification was performed using the external standard method with calibration curves ranging from 0.01 to 6 mg mL⁻¹, and the results were presented as percentages of each lipid class.

2.2.4. Total Dietary Fiber Content

The dietary fiber (Tdf) content was determined following AOAC Mes-Tris method 991.43 [19]. This procedure utilized three enzymes (thermostable α-amylase, protease, and amyloglucosidase) under specific incubation conditions to break down starch and protein components. Non-digestible residues, representing dietary fiber fractions, were obtained after enzymatic digestion. Insoluble dietary fiber (Idf) was separated by filtration, while soluble dietary fiber (Sdf) was precipitated using ethanol. The dry residues corresponded to Idf and Sdf, and the total dietary fiber (Tdf) was calculated as their sum. The yield of dietary fiber (df) was determined using the equation

df yield (%) = DF × 100/W

(1)

where df is the content (g) of Tdf, Idf, or Sdf as appropriate and W is the weight (g) of the sample.

2.2.5. Total Phenolic Content

The total phenolic content (TPC) was determined by the Folin–Ciocalteu spectrophotometric method at 760 nm, using hydroxytyrosol (HT) as standard within the range of a 0–1 mg L⁻¹ calibration curve [20]. TPCs were given as mg of equivalent HT per kg of oil (mg eHT/kg). These determinations were carried out in duplicate.

2.2.6. Phenolic Compound Characterization by UHPLC-ESI-MS

Sample preparation was carried out according to Aturki et al. [21]. The PCs were extracted with 5 mL MeOH:H₂O (80:20, v/v) twice from 1 g of DF. The methanolic phase separated by centrifugation (at 4500 rpm for 10 min) was evaporated, to obtain the phenolic extract, at 40 °C and under vacuum (~1 mbar) using a rotary evaporator, RV 10 (IKA^®-Werke GmbH & Co. KG, Staufen, Germany), and an oil-sealed rotary vacuum pump vacuum (Edwards, Spain distributor Iberica Vacuum, Madrid, Spain). The PCs were analyzed by ultrahigh-performance liquid chromatography (UHPLC) coupled to an EvoQ Elite (Bruker, Billerica, MA, USA) mass spectrometer equipped with an electrospray ionization (ESI) source in the MS/MS mode. The analysis was carried out at the Chromatography Unit of the Interdepartmental Research Service of the Autonomous University of Madrid (SIDI-UAM) according to [22]. Samples were stored at 4 °C before injecting 5 µL in the full-loop mode onto an ACE Excel 3 C18 column (3 × 150 mm, 3 µm, Avantor, Radnor Township, PA, USA). Chromatographic separation was carried out using a gradient of solvent A (H₂O with 0.1% acetic acid) and solvent B (acetonitrile with 0.1% acetic acid) at a flow rate of 0.4 mL min⁻¹. The gradient profile was as follows: 10% B for 5 min, 50% B at 10 min, and 100% B at 15 min (maintained for 2 min), followed by a return to 10% B in 0.2 min, giving a total run time of 20 min. The column temperature was maintained at 21 °C. Nitrogen was used as cone gas (40 L h⁻¹), heated probe gas (50 L h⁻¹), and fogging gas (60 L h⁻¹). The cone and probe temperatures were set at 350 °C and 400 °C, respectively, with a capillary voltage of 4.5 kV. Mass spectra and tandem mass spectra were acquired in the negative ion electrospray ionization (ESI) mode, with a quadrupole mass analyzer set to a −1 m/z precursor ion selection window. The mass spectrometer was calibrated in the negative ion electrospray ionization (ESI) mode. The mass spectrometer was calibrated using an external standard mixture. Samples were dissolved in MeOH:H₂O (80:20), and calibration curves were prepared for quantifying hydroxytyrosol, gallic acid, tyrosol, vanillic acid, luteolin-7-o-glucoside, isoquercitrin, ferulic acid, p-coumaric acid, elenolic acid, luteolin, pinoresinol, erythrodiol, apigenin, caffeic acid, syringic acid, and oleuropein. All samples were extracted and analyzed in duplicate.

2.2.7. Aqueous Extract of HD

A double solid–liquid extraction of the defatted flour (DF) was carried out with water at a ratio of 1:10 (DF:H₂O). The DF mixture was left for 10 min in an ultrasonic bath, then homogenized with Ultra-turrax T25 (IKA, Tucson, AZ, USA) at 10,000 rpm for 2 min and was centrifuged at 3500 rpm for 20 min, at room temperature [9]. The aqueous phases were collected and used for the preparation of films.

2.3. Film Elaboration

The active film solutions were prepared according to the method described by Vidal et al. [9], with slight modifications. A total of 2 g CMC was mixed with 70 mL ultrapure water in an Ultra-turrax T25 (IKA, Tucson, AZ, USA) for 2 min, then taken to a magnetic stirrer for 30 min (70 °C, 600 rpm). Separately, 1 g of DF was dissolved in 30 mL of ultrapure water for 5 min at 25 °C, also using Ultra-turrax. Then, the solutions were mixed to obtain a DF/CMC ratio of 1:2, 0.5 mL glycerol was added as a plasticizer (25 wt.% CMC) and 0.08 g phosphatidylcholine (PC) as an emulsifier (4 wt.% CMC), and the mixture was homogenized with the Ultra-turrax at 13,500 rpm for 10 min.

For the preparation of DFE films, the same procedure was carried out with 80 mL of ultrapure water to dissolve 2 g of CMC, and the 20 mL of the aqueous phase obtained in the previous section, Section 2.2.7, was added. A pure CMC control film was prepared by dissolving 2 g of CMC in 100 mL of ultrapure water and adding glycerol and PC in the same proportions. In total, 25 mL of the resulting solution was poured into polystyrene Petri dishes (90 mm diameter) and dried for 24 h at 25 °C in a fume hood. The dried films were manually peeled off the plates and stored in a desiccator at 58% relative humidity (RH) until use.

2.4. Characterization of the Obtained Films

2.4.1. Weight and Film Thickness

The processed films were weighed and their thickness was measured with a micrometer (Jeankak, Shenzhen, China; accuracy: 0.001 mm). Thickness measurements were taken at 10 random points on each film and mean values were calculated to determine the weight and thickness of the films.

2.4.2. Film Moisture Content and Water Solubility

The films were cut into small pieces (500 mg, w₁) and dried in a porcelain container at 105 °C to constant weight (w₂). The film moisture content (%) was obtained by Equation (2):

Moisture (%) = ((w₁ − w₂)/w₁) × 100

(2)

After obtaining w₂, the film pieces were immersed in 50 mL of distilled water and left to soak for 24 h at room temperature. The wet samples were then dried at 105 °C for 24 h to determine their final dry mass (w₃). The water solubility of the films was calculated according to the following equation, Equation (3):

Solubility (%) = ((w₂ − w₃)/w₂) × 100

(3)

2.4.3. Transparency and Film Transmission

The UV-Vis barrier properties of the films were assessed by exposing freshly prepared film strips (10 × 40 mm) to wavelengths ranging from 200 to 800 nm. An empty test cuvette served as the reference. The transparency (T) value of the films was calculated using the following equation, Equation (4):

T = −log(t₆₀₀)/Δx

(4)

where t₆₀₀ is the transmittance at 600 nm and Δx is the film thickness (mm). Three replicates were performed for each film formulation.

2.4.4. Film Total Phenolic Content

TPC of the films was determined by the Folin–Ciocalteu spectrophotometric method using the same parameters as in Section 2.2.5. These determinations were performed in triplicate.

2.5. Bioactive Packaging Films to Prevent Lipid Oxidation

Commercial microalgae oil with high DHA content (50%) from the marine microalga Schizochytrium sp. was used to evaluate the oxidation protection of the films produced. In 80 cm Petri dishes, 25 mL of oil was placed and capped with the films (CMC, CMC + DF, CMC + DFE). The headspace was purged with N₂ to remove internal residual O₂ at the beginning of the experiments and after each sampling. An unprotected control blank was also prepared. Samples were kept on an orbital shaker at 40 °C and 110 rpm for 10 days.

The oxidation state of the oil was evaluated by measuring the peroxide value (PV), expressed as milliequivalents of active oxygen per kilogram of oil (meq O₂ kg⁻¹), which served as an indicator of primary oxidation. PV was determined using 5 μL of oil by colorimetric reactions measured with an Oxitester (CDR FoodLabFat, Florence, Italy), following the manufacturer’s protocol according to the official AOCS Cd 8–53 methods [23]. Absorbance readings were performed at 505 nm. All measurements were performed in duplicate.

2.6. Statistical Analysis

A data analysis was performed using Excel (Microsoft Office) and all statistical evaluations were performed using Origin (version 9.0 for Windows; OriginLab Corporation, Northampton, MA, USA). The data were expressed as the mean ± standard deviation. The statistical significance of the differences between the groups was measured by a one-way analysis of variance (ANOVA) and post hoc Tukey HSD test. Statistical significance was defined at the level of p < 0.05.

3. Results and Discussion

3.1. Physicochemical Parameters of Olive Defatted Flour

Table 1. Physicochemical parameters of olive defatted flour obtained during supercritical fluid extraction. Humidity (%), Granulometry (%), TPC (total phenolic content (ppm)), TFC (total fat content (%)), lipid composition, and total dietary fiber (%) (IDF: Insoluble dietary fiber and SDF: Soluble dietary fiber).

Olive Defatted Flour
Humidity (%)	2.17 ± 0.33
Granulometry (%)
Diameter (µm)
>500	32.31 ± 1.23
250–500	27.81 ± 0.50
100–250	24.70 ± 1.12
45–100	13.93 ± 0.63
<45	1.37 ± 0.22
TFC (%)	2.43 ± 0.21
Lipid composition (%)
Fatty acid	0.71 ± 0.19
DAG	1.74 ± 0.14
Sterol esters	1.58 ± 0.28
TAG	42.72 ± 9.69
TPC (ppm)	34,856 ± 1220
Total dietary fiber (%)
IDF	28.40 ± 1.75
SDF	2.28 ± 0.34
IDF:SDF	12.62 ± 1.11

presents the physicochemical properties of the DF used in this study. The moisture content is 2.17%. This value is lower than the 8–10% obtained in the olive cakes after oil extraction with organic solvents [24,25]. Regarding the particle size parameter of the flour, it is below 500 µm. This is due to the shearing effect produced by the agitation during oil extraction, which improves yields and allows us to obtain flours whose size does not need to be reduced to produce films [6].

Regarding the total fat content (TFC) of the DF, it is 2.4%, similar to the content of the hexane defatted olive cakes (1.53 ± 0.4%) [25]. Approximately 50% of the TFC are neutral lipids, of which the majority are triglycerides. The other half could be polar lipids and phenolic compounds of an apolar nature, since the TPC is ≈35,000 mg eHT per kg of DF. These values are similar to those reported by Zhao et al. where dehydrated and defatted olive pomace was analyzed with hexane and the values are 36,000 ppm [26]. Despite this, the DF obtained after SFE is better for incorporation into food formulations due to the absence of any solvent residue [27].

Total dietary fiber (Tdf) levels (Table 1) of the DF are remarkable and similar to those found in olive pomace [25,28]. Likewise, insoluble dietary fiber (Idf) is the main fraction of Tdf, as it accounts for more than 90% of Tdf; thus, the Idf/Tdf ratio is very high (12.6). Idf mainly favors digestive health by accelerating intestinal transit due to it being characterized by not dissolving in water or forming gels, which has less effect on blood glucose and cholesterol control, but provides a greater feeling of satiety [29]. High Idf can improve stiffness and reduce water sensitivity in a packaging film, but will also require formulation modifications to balance its structure and avoid excessive stiffness or brittleness [30].

3.2. The Characterization of the Phenolic Fraction from the Olive Defatted Flour

Total phenolic compounds were characterized by UHPLC-ESI-MS under the aforementioned conditions, searching only for compounds whose standards were available in the laboratory (Figure 1). A total of 15 peaks were identified and quantified in the chromatograms, and the PC content is shown in

Table 2. Assignment and content of phenolic compounds of defatted flour (DF), mean ± standard deviation expressed in mg/kg of DF, retention time (Rt), and deprotonated molecule ([M − H]⁻). n.q.: not quantified.

Peak No.	Class/Phenolic Compounds	Rt	[M − H]− (m/z)	mg/kg DF
1	Gallic acid	2.78 ± 0.00	169	1.69 ± 0.33
2	Hydroxytyrosol (HT)	3.69 ± 0.02	153	1524.74 ± 178.16
3	Tyrosol (TYR)	4.92 ± 0.57	137	156.86 ± 41.95
4	Caffeic acid	4.63 ± 0.00	179	0.76 ± 0.12
5	Syringic acid	4.66 ± 0.01	197	0.25 ± 0.03
6	Vanillic acid	4.72 ± 0.00	167	1.40 ± 0.17
7	Luteolin-7-O-glucoside	4.92 ± 0.00	447	210.56 ± 10.10
8	Isoquercitine	4.93 ± 0.00	463	2.27 ± 0.39
9	Ferulic acid	5.50 ± 0.00	193	1.02 ± 0.00
10	Oleuropein	5.53 ± 0.01	539	4324.74 ± 408.87
11	p-Coumaric acid	5.66 ± 0.02	163	0.05 ± 0.00
12	Elenolic acid (EA)	6.08 ± 0.02	241	3602.75 ± 1555.62
13	Luteolin	6.47 ± 0.00	285	13.31 ± 0.68
14	Pinoresinol	6.66 ± 0.00	357	9.22 ± 0.63
15	Erythrodiol	6.81 ± 0.02	441	n.q.
16	Apigenin	7.08 ± 0.00	269	0.45 ± 0.02
Σ Polyphenols				8891.11 ± 763.72

. According to Table 1, the compounds identified in Table 2 represent 30% of the total phenols present in the DF. The phenolic alcohols identified were hydroxytyrosol (HT, peak 2) and tyrosol (TYR, peak 3). Caffeic acid (peak 4), syringic acid (peak 5), vanillic acid (peak 6), and ferulic acid (peak 9) were identified as phenolic acids. As for secoiridoids, the peaks identified were oleuropein (peak 10) and its secondary metabolite, elenolic acid (peak 12). Flavonoids were also identified, such as luteolin-7-O-glucoside (peak 7), luteolin (peak 13), and apigenin (peak 16), and the lignan pinoresinol (peak 14). Other studies show that all these compounds are found in olive pomace [31].

The most abundant phenolic compound in DF is oleuropein (4324 ppm), whose content is much higher than that usually found in olive pomace (2600 ppm) [32]. This is mainly due to the fact that oleuropein is degraded during the crushing and malaxation of olives in the presence of water because its glycosidic bond is easily hydrolyzed, producing oleuropein aglycone, whose secoiridoid ring opens, giving rise to other derivatives such as elenolic acid [32,33]. Pharmacological research on this oleuropein derivative is of great importance as it exhibits promising preventive and therapeutic potential [34]. Elenolic acid along with HT, luteolin-7-O-glucoside, and TYR are the other major PCs in DF with amounts of 3603, 1525, 211, and 157 ppm, respectively.

As mentioned above, DF is a valuable source of bioactive substances that offer established benefits for human health and well-being. In particular, it is rich in phenolic compounds, which have been associated with various biological activities, such as anti-inflammatory, antitumor, antimicrobial, antioxidant, antidiabetic, and cardioprotective effects [35]. Among these bioactive phenols, HT stands out as one of the most valuable compounds recoverable from the solid by-product, due to its high oxidative stability and potent antioxidant properties [31]. HT is currently used as a therapeutic agent, dietary supplement, and natural ingredient both as a biological and technological antioxidant [36]. Given the enormous amount of bioactive phenolic compounds present in DF together with its high insoluble fiber content and the fact that it can be considered a food-grade ingredient, since no organic solvents are used in the whole procedure, it could potentially be used as an additive to bakery products or other food matrixes. Alternatively, it could be incorporated in food packaging, providing food with a bioactive effect and an environmentally sustainable option.

3.3. Film Characterization

3.3.1. Film Weight, Thickness, Moisture Content, and Water Solubility

Table 3. Physical properties and water solubility (WS) of control film (CMC), film with defatted flour (CMC + DF), and film with DF extract (CMC + DFE).

Films	Weight (g)	Thickness (µm)	Moisture (%)	WS (%)
CMC	0.68 ± 0.05 b	62 ± 7.49 c	6.8 ± 0.25 c	100 a
CMC + DF	0.82 ± 0.02 a	108 ± 16 a	12.5 ± 1.51 a	52.6 ± 2.62 b
CMC + DFE	0.72 ± 0.03 b	80 ± 8.10 b	9.7 ± 0.73 b	100 a

Significant differences (p < 0.05) are indicated in the same column with different letters (a, b or c).

shows the weight, thickness, moisture, and water solubility of the films produced. By adding the flour (CMC + DF), the weight, thickness, and moisture content of the film increase significatively (p < 0.05) with respect to the control film (CMC) and the film containing the extract (CMC + DFE). The shelf life of food products depends on the transfer of water between the product and its surrounding environment. To preserve freshness, films should minimize this water transfer. Moisture results are lower than those obtained by de Moraes Crizel et al. [37] by adding up to 30% olive pomace to films prepared with chitosan (12.5 vs. 16.5%). Although the film with DF has less water solubility than the control and DFE films, this is probably due to the insoluble fiber in the flour and the presence of hydrophobic compounds since it contains 2.4% TFC [9,37]. The high solubilities make these films more suitable for dry or low-water-activity products. In future studies, it is suggested to modify the formulation by incorporating hydrophobic agents such as oil or waxes for use in wet products.

3.3.2. Light Transmission and Transparency

Optical characteristics are critical for determining the effectiveness of packaging as a food preservation agent when exposed to visible and ultraviolet light [38]. The higher the transmittance at a given wavelength, the lower the absorbance of the light, and thus the lower the protection of the film [39]. As can be seen in

Table 4. Light transmission and transparency of control film (CMC), film with defatted flour (CMC + DF), and film with DF extract (CMC + DFE).

Films	Transmittance (%)						Transparency (%)
	200 nm	300 nm	400 nm	500 nm	600 nm	800 nm	600 nm
CMC	0.07 ± 0.01 a	67.3 ± 0.04 a	82.8 ± 0.00 a	85.5 ± 0.01 a	86.7 ± 0.00 a	88.2 ± 0.01 a	1.03 ± 0.03 c
CMC + DF	0.02 ± 0.01 b	0.02 ± 0.00 b	0.00 ± 0.00 c	0.71 ± 0.00 c	3.72 ± 0.00 c	14.2 ± 0.00 c	11.5 ± 1.70 a
CMC + DFE	0.03 ± 0.01 b	0.02 ± 0.00 b	1.23 ± 0.01 b	19.6 ± 0.01 b	38.9 ± 0.01 b	64.7 ± 0.00 b	5.0 ± 0.15 b

Significant differences (p < 0.05) are indicated in the same column with different letters (a, b or c).

, the transmittance increases as the wavelength increases. At the UV wavelengths of 200 and 300 nm, the films incorporated with DF and DFE demonstrated better UV barrier properties, while the control film exhibited a light transmission of 67.3% at the 300 nm wavelength, compared to 0.02% for DF and DFE. At a higher-visibility spectrum (400–800 nm), DF reduced between 1.2 and 27 times the visible light transmission absorbance capacity compared to DFE films. These results agree with those obtained in [9] for CMC films with green coffee sediment, and those obtained by de Moraes et al. (2018) [37] for chitosan films, in which they add up to 30% olive pomace. The transparency values of the films added with DF and DFE were significantly higher than those of the control film, the films with DF being the ones with higher opacity and consequently more efficient in light barriers [40]. This occurs because DFE solubilizes better in the film matrix, resulting in a more homogeneous matrix as shown in Figure 2. Therefore, a higher protection is expected from DF film [37]. However, these preliminary studies need to be corroborated with others to assess the impact of prolonged exposure under real conditions.

3.3.3. Total Phenolic Content of the Films

Figure 3 shows the values of total phenolic content (TPC) expressed in mg of equivalents HT per kilogram (mg eHT/kg), in DF, CMC, CMC + DF, and a mixture of CMC with an aqueous extract of DF (CMC + DFE). As mentioned above, DF has the highest content of phenolic compounds, which underlines its potential as a rich source of antioxidants. When the flour or its aqueous extract is combined with CMC, the phenol content decreases considerably, possibly due to dilution or an interaction between the flour compounds and the CMC matrix. Another factor could be the solvent used in the Folin–Ciocalteu method to extract phenolic compounds from the films, since they are extracted with methanol/H₂O (60:40, v/v). In water alone, the hydrophilic nature of CMC promotes polymeric chain relaxation, causing substantial swelling that dissolves most of the active films, thereby facilitating the complete release of phenolic compounds [41].

3.4. Lipid Oxidation Control Using Bioactive Packaging Films

Figure 4 shows the peroxide (PV) values of the DHA-rich microalgae oil at 40 °C under agitation. For the uncapped control oil, the PV began to increase significantly from the first day, as expected, reaching 50 meqO₂/kg in 5 days. These values agree with those obtained in a previous study of oxidative stability of microalgae oil [42]. In the case of the oil covered with CMC film, its PV began to rise drastically after the fourth day, reaching values close to 50 meq O₂/kg in 6 days. Protective films significantly (p < 0.05) reduce peroxide formation rates, with CMC + DF film standing out as the most effective in minimizing oxidation, followed closely by CMC + DFE (9 and 8 days, respectively).

The regulation sets the primary oxidation limit (peroxide value) for oils rich in LCPUFAs at 5 [43]. With the control film, it takes 1–2 days to reach this value, while in the presence of the films with DF and DFE, the oils take 4–5 days to reach this value under the accelerated conditions (40 °C, 110 rpm). This represents a protection of more than 100% against the control film. This superior performance is due to the high concentration of antioxidant compounds, including phenolics from olive residues, which actively combat oxidative reactions. In addition, the natural opacity of the films provides a protective barrier against photooxidation, making them ideal for preserving product quality and extending shelf life [38]. These results agree with those found in the literature, since there are studies on the incorporation of olive pomace flours in packaging films, olive leaf extracts, grape pomace, or coffee by-product extracts. In the literature, the lipid oxidation of different foods has been studied, and it has been found that the presence of antioxidant compounds in packaging, in addition to its barrier against water, oxygen, and light, positively prevents lipid oxidation [9,37,39,41,44]. However, these are preliminary data to evaluate whether the film could reduce the oxidation rate of a polyunsaturated oil such as microalgae oil, for a possible use as an encapsulant of these omega-3 oils. Given the potential of the films, further studies at different temperatures are needed to extrapolate the results at room temperature and study the oxidative stability of the oil.

Finally, both DF and DFE maintain their food-grade quality throughout the manufacturing process, ensuring that they are suitable for direct contact with food products. This makes them useful not just as protective food packaging, but also as potential transporters of bioactive oils, which may improve the nutritional and functional qualities of packed foods. Furthermore, the antioxidant-rich composition of DF and DFE films adds an extra layer of protection against oxidative degradation, improving the shelf life and quality of the encapsulated bioactive components. As a result, potential applications of these films could be the encapsulation of oils rich in polyunsaturated fatty acids, as indicated above. However, digestion studies are needed to determine the bioaccessibility of the oils and bioactive compounds in the capsule. In addition, other possible uses would be internal coatings for boxes, bags for dry products, or wrappings for small foods. Finally, the formula could be adapted for wet products, incorporating hydrophobic compounds.

4. Conclusions

The defatted flour (DF) showed a significant content of oleuropein, elenolic acid, and hydroxytyrosol, among other phenolic compounds. The oxidation of the oil protected with CMC + DF and CMC + DFE film was slower than that of the oil protected only with CMC film. The use of DF increases the UV-Vis barrier capacity and antioxidant capacity, which means that for foods rich in polyunsaturated fatty acids that require higher light protection, CMC + DF films are more suitable. Both DF and DFE preserve food grade throughout the manufacturing process, so the films could be used to encapsulate bioactive oils as well as food packaging. Nevertheless, the main focus of this study was the initial characterization of defatted flours from a clean olive oil extraction process, as well as their possible use for bioactive packaging by studying the antioxidant and physicochemical properties of the films. Future work is also suggested to include mechanical properties and phenolic compound migration tests to ensure the safety and functionality of the material.