Microencapsulation of Chia Oil Using Whey Protein and Gum Arabic for Oxidation Prevention: A Comparative Study of Spray-Drying and Freeze-Drying Methods

Abstract

Encapsulation is one of the most widely utilized strategies for preventing oil oxidation. Chia oil is a source of oils that are high in omega-3s and was used as a core material in this study. Whey protein and gum arabic were used as wall materials, and the whey protein:gum arabic ratios were 1:0, 1:1, 1:2, 2:1, 1:3 and 3:1. Preparation of chia oil microcapsules was conducted by spray-drying or freeze-drying methods. All microcapsules were stored in an opaque and airtight laminate pouch for 3 and 6 months to determine the effect of time on the fatty acid composition, encapsulation efficiency and chemical properties. Encapsulation had a positive protective effect on oil quality. The two drying methods resulted in different powder characteristics: spray drying resulted in a spherical shape, whilst freeze drying led to flakes and a porous surface. Spray drying microcapsules resulted in greater encapsulation efficiency than freeze drying microcapsules. In addition, encapsulated powders stored for 6 months showed both acid and peroxide values below the Codex limit. These results indicate a possible encapsulation process to protect chia oil from oxidation.

1. Introduction

Current food trends indicate a substantial increase in the consumer demand for food with higher nutritional value, promoting health and enhancing the body’s physiological functions. The role of fatty acids in current health and diets has sparked particular interest in the consumption of polyunsaturated fatty acids, which include omega-3, omega-6 and omega-9 polyunsaturated fatty acids [1]. Among vegetable sources, chia (Salvia hispanica L.) is indigenous to Southern Mexico and has been cultivated and is an essential part of Mesoamerican cultures and diets [2]. Chia seeds contain between 28 and 32% oil, and chia oil is one of the best sources of omega-3 fatty acids (61 to 70%). Daily ingestion of 1 g of chia oil satisfies the minimum requirements for omega-3 fatty acids in adults [1]. Increased omega-3 oil consumption can also be achieved by fortifying foods such bread, milk and yogurt with omega-3 fatty acids [3]. Omega-3 (ω-3) fatty acids are nutritionally essential for overall health and can help those with diabetes, coronary heart disease and immune response issues [4,5]. Indeed, the addition of omega-3 fatty acids to food is limited due to their oxidative instability and the formation of oxidized compounds, resulting in a shorter shelf-life of the food. Therefore, the microencapsulation of polyunsaturated fatty acids comprising omega-3- and omega-6-rich oils is presently regarded as a promising and interesting food science technological option [6]. It has previously been reported that microencapsulation can improve the stability of chia oil [7].

Microencapsulation is the technique of enclosing small particles or droplets by a coating wall in a homogeneous or heterogeneous matrix to provide a barrier between the component inside the capsule and its surroundings [8]. This technology is being extensively studied because it can prevent oil oxidation and minimize off-flavors in some food items, hence preserving their nutritional value [9,10,11]. This process involves emulsifying a core material, which is usually a lipid, with a dense solution of wall material, such as proteins, gums or carbohydrates. The emulsion is then atomized and dried [7].

From a technological point of view, the combination of lipophilic substances, such as oils, has a restricted ability to be incorporated into various forms of food matrices, particularly in water-based carriers due to their poor solubility and hydrophobicity, as these molecules are insoluble. The formation of hydrophilic powders incorporating oil provides a potential solution to this issue.

Carbohydrates and proteins are commonly used for the microencapsulation preparation of ω-3 fatty acids and oils. These compounds can be combined with other components, such as antioxidants, surfactants, oxygen scavengers and chelating agents, to achieve specific functions in food or nutraceutical applications [12]. In addition, various protein/polysaccharide combinations have been described in the literature as potential wall materials, including chitosan, maltodextrins, dextrins, modified starches, agar, starches, alginates, carrageenan, pectin, gum acacia, gum arabic and mesquite gum [13,14]. Among the wall materials used for microcapsule encapsulation, polysaccharides have been one of the most used wall material compounds [15]. Gum arabic (GA) is recognized as an effective emulsifier, possessing desirable attributes such as being colorless, odorless, tasteless, non-toxic and non-irritating. It also has low viscosity and provides excellent protection against oxidation, making it suitable for use as a filler or drying agent in various applications. GA is a type of complex heteropoly electrolyte that is characterized by highly branched structures primarily composed of L-arabinose and D-galactose, as well as smaller amounts of 4-O-methyl-D-glucuronate and L-rhamnose [16,17]. However, the protein content of GA is relatively low, ranging from 1.0% to 2.0%, which is responsible for its emulsifying and film-forming properties [18]. Proteins have become increasingly popular as wall materials, particularly in the fields of nutrition, cosmetics and pharmaceuticals. This is due to their excellent surface-active properties, which make them well suited for microencapsulation that requires prior emulsification. Gelatin, caseinates, gluten, albumin, vegetable proteins, soy protein, zein and whey protein (WP) are among the most commonly used proteins for microencapsulation. WP is composed of a diverse range of proteins that exhibit functionality in a broad range of food applications. WP has a globular structure that remains stable in solutions across a wide pH range, but it can be denatured by heat and exhibits heat-induced gelation [19].

The encapsulation process utilizes various techniques that involve creating droplets of the active compounds and enveloping them with a carrier material through various physicochemical processes in a gas or liquid phase [20]. The most commonly utilized and well-established processes for encapsulation are spray drying (SD) and freeze drying (FD).

Microencapsulation by spray drying is extensively employed in the food industry for the formulation of some dry extracts, such as flavors and essential oils [21,22]. Spray drying (SD) involves spraying a liquid product such as an emulsion, suspension or solution into a hot gas to evaporate the solvent and produce a powder. However, the high drying temperatures used in SD can pose a risk for microcapsules that contain polyunsaturated oils. On the other hand, freeze drying (FD) involves freezing the product and removing water through sublimation, which reduces structural degradation. Additionally, the use of very low temperatures and the absence of air in FD help to prevent product deterioration.

Currently, there is a lack of literature describing the properties of products resulting from the microencapsulation of chia oil using freeze-drying (FD) and spray-drying (SD) processes. The objective of this study is to formulate and document the microencapsulation of chia oil using whey protein and gum arabic as wall materials through the SD and FD processes in order to produce long-lasting powders. The study includes a comparative analysis of the efficacy of the processes and the physicochemical and morphological characteristics of the microcapsules. Additionally, the study examines the oxidative stability of the microcapsules with varying storage times.

2. Materials and Methods

2.1. Materials

Chia seeds were provided by the Plant Breeding Center for Sustainable Agriculture, Faculty of Agriculture, Khon Kaen University Khon Kaen, Thailand. Chia oil was extracted from the seeds using hexane (at a ratio of 1 g chia seed to 5 mL hexane) for 12 h at room temperature and 200 rpm. The oil was then filtered using Whatman filter paper grade 1, and the filtered oil was stored in amber bottles under N₂ gas at approximately 4 °C until needed.

The following chemicals were used as wall materials/emulsifiers in this work: whey protein (WP) was purchased from the market (Proflex brand); WP was produced by Siamcanery (Ban Bueng, Chonburi, Thailand); gum arabic (GA) was purchased from Union Science Co., Ltd., Chiangmai, Thailand. Double-distilled water was used in all experiments. Hexane, heptane, dichloromethane, petroleum ether, chloroform, acetic acid, sodium hydroxide, potassium iodide and sodium thiosulfate were bought from Sigma Aldrich, Singapore. Starch was purchased from a local distributor from Mengseng Starch Co., Ltd. (Nakhonratchasima, Thailand).

2.2. Proximate Analysis and Fatty Acid Determination

The proximate composition of the chia seeds was obtained by the standard method of the Association of Official Analytical Chemists [23]. Moisture content was assessed by drying the samples in an oven at 105 °C until a constant weight was obtained. Ash content was determined by incinerating known weights of the samples in a muffle furnace at 550 °C (Method No. 930.05). The Soxhlet extraction method (Method No. 930.09) was used to extract crude fat by exhausting a known weight of the sample in petroleum ether. Crude protein content (N × 6.25) was measured by the Kjeldahl method (Method No. 984.13). Crude fiber quantity was determined by digesting a known weight of the fat-free sample in refluxing 1.25% sulfuric acid and 1.25% sodium hydroxide (Method No. 962.09). Carbohydrates were calculated by subtracting the sum of the other components from 100. All experiments were carried out in triplicate.

The methylation reaction was performed before fatty acid analysis by gas chromatography (GC). The reaction was performed by mixing 500 μg of chia oil with 1 mL mixed solution of methanol:toluene:2,2-dimethoxypropane:H₂SO₄ (60:20:5:2, v/v) [24] and 500 μL heptane and then incubated at 80 °C for 2 h. GC was used to analyze the fatty acid composition of the chia oil using an Agilent Technologies GC7890 instrument with an autosampler (model 7693), a flame ionization detector (FID) and a Stabilwax^®-MS column (30 m × 250 µm × 0.25 µm). The analysis started at an initial column temperature of 210 °C with a hold time of 5 min, followed by a heating ramp of 20 °C/min until 230 °C was reached with a hold time of 4 min. Helium was used as a carrier gas at a flow rate of 1 mL/min.

2.3. Emulsion Preparation

To adjust the protein/polysaccharide ratio of the microcapsule wall, six samples were prepared using WP 100% and WP/GA at ratios of 1:1, 1:2, 2:1, 1:3 and 3:1, which were then dried using the SD method. These samples were labeled as W10S, A11S, B12S, C21S, D13S and E31S, respectively. Similarly, six different samples were prepared using a similar wall material composition as previously prepared, but were dried by the FD method (sample names W10F, A11F, B12F, C21F, D13F and E31F, respectively). The ingredients listed in

Table 1. Composition of the emulsion containing different whey protein to gum arabic ratio.

Code	Weight of Core Material (g)	Weight of Wall Material (g)			WP:GA Ratio	Core to Wall Material Ratio	DDW (mL)
	Chia Oil	WP	GA	Total
W10	14.68	73.72	0	73.72	1:0	1:5	500
A11	14.68	36.86	36.86	73.72	1:1	1:5	500
B12	14.68	24.57	49.15	73.72	1:2	1:5	500
C21	14.68	49.15	24.57	73.72	2:1	1:5	500
D13	14.68	18.43	55.29	73.72	1:3	1:5	500
E31	14.68	55.29	18.43	73.72	3:1	1:5	500

WP: whey protein, GA: gum arabic, DDW: double-distillated water.

were used to prepare the emulsions in all cases. WP and GA were dissolved in 500 mL of double-distilled water, mixed thoroughly using magnetic agitation for 30 min and kept under magnetic stirring at room temperature until completely dissolved. Next, 14.68 mL of chia oil was added dropwise to each emulsion, while stirring at 100 rpm for 12 h at room temperature to ensure complete protein hydration. The resulting emulsions, with a volume of approximately 550 mL, were stored at 4 °C before starting the drying process.

2.4. Spray-Drying Process

The laboratory-scale spray-drying process was carried out using a Buchi Mini Spray Dryer B-290, equipped with a double fluid type atomizer nozzle and a 0.7 mm spray mesh, in a drying chamber with dimensions of 16.5 cm × 60 cm (H × D). The peristaltic pump was operated at a flow rate of 5.5 mL/min, and the air outlet temperature was carefully maintained at 82 ± 3 °C to optimize the final characteristics of the encapsulated powder. The drying air flow rate, air pressure and compressed air flow rate were 35 m³/h, 6 bars and 1 m³/h, respectively, while the drying air inlet temperature was fixed at 150 °C [25]. The equipment dehumidifier, model R-296 (Buchi Labortechnik AG, Flawil, Switzerland), was used to provide air with similar relative humidity for all drying trials (40%). The resulting microparticles were kept in a desiccator at room temperature.

2.5. Freeze-Drying Process

The emulsion was frozen at −80 °C for 12 h in round bottle flasks before being connected to the freeze-dryer unit. The freeze-drying (FD) process was carried out using a laboratory FreeZone^® Console Freeze Dryers (Labconco, Kansas City, MO, USA) under a vacuum of 0.1 mbar and with a condenser set at −80 °C for 48 h. The round bottle flasks were maintained at room temperature (25 ± 2 °C) during the FD process. Microparticles produced by freeze drying were also kept in a desiccator at room temperature.

2.6. Recovered Solid Yield

The recovered solid yield (SY) was determined by dividing the weight of the final powder obtained after each drying experiment (W_fi, dry basis) by the initial weight of the components used in the emulsion’s preparation (excluding water) (W_in, dry basis). The calculation is expressed in Equation (1).

SY = (W_fi/W_in) × 100

(1)

where W_fi is the weight of powder collected after the following drying experiment and W_in is the weight of the initial number of factors in the emulsion (except water).

2.7. Color Measurement

A colorimeter (Mini Scan E2 Manual Version 1.2) was used to measure the color of the samples with D65 (daylight) and an inclination of 10°. The color values of the samples were expressed in terms of L* (lightness), a* (redness/greenness) and b* (yellowness/blueness) parameters.

2.8. Moisture Content and Water Activity

The moisture content of the encapsulated powders was determined by drying in a forced-circulation oven at 105 ± 1 °C until a constant weight was achieved, and the gravimetric method was used. The water activity meter (Aqualab series 3; Decagon Devices Inc., Washington, DC, USA) was calibrated with saturated LiCl before determining water activity. All measurements were taken at room temperature during the test, and all determinations were performed in triplicate immediately after drying [25].

2.9. Morphological Analysis

The microstructural properties of the SD and FD encapsulated powders were investigated using a scanning electron microscope (LEO-1450VP model, Carl Zeiss—Sigma, Oberkochen, Germany). The samples were mounted on SEM stubs with two-sided adhesive tape (Ted Pella, Redding, CA, USA), and coated with gold using a magnetron sputter coater (Denton Vacuum Model, Moorestown, NJ, USA) at 100 millitorrs and 15 mA. The coated samples were then analyzed using the SEM operating at an accelerating voltage of 25 kV. Micrographs depicting the microstructure of the microencapsulated powder were obtained using the instrument’s software installed on a PC connected to the SEM.

2.10. Total Oil Determination

The determination of total oil (TO) in the microencapsulated powder was carried out with slight modifications to a previously reported method. Briefly, 1.0 ± 0.1 g of each sample was placed in a Soxhlet extraction apparatus along with 50 mL of n-hexane and extracted for 3 h. The solvent was then evaporated, and the remaining oil was heated at 60 °C until a constant weight was achieved. The weight of the extracted oil was measured and expressed as a percentage of the microcapsule weight (dry basis). This procedure was performed in triplicate for each sample.

2.11. Determination of Encapsulation Efficiency

The method used to determine the surface oil content (SO) of microcapsules was based on a previous study with some modifications [26]. Initially, 1.00 ± 0.01 g of the sample was weighed and mixed with 20 mL of petroleum ether. The resulting mixture was stirred for 1 min and filtered. The solids obtained were washed with 10 mL of petroleum ether, and the organic phases were combined. The solvent was then evaporated, and the remaining oil was heated at 60 °C in an oven until a constant weight was achieved. This experiment was carried out in triplicate. The percentage of encapsulated oil (%EE) was determined by calculating the ratio of the total oil contained in the microcapsules and the oil located on their surface (SO) using Equation (2).

%EE = [(TO − SO)/TO] × 100

(2)

where %EE is the percentage of encapsulation efficiency, TO is the total oil extracted from the microcapsule and SO is the surface oil content.

2.12. Analysis of Oil’s Oxidative Stability at Different Storage Times

All the encapsulated powders were stored in vacuum-sealed aluminum pouches at room temperature (25–30 °C) for up to 6 months. The peroxide value (PV) of the samples was examined at different intervals (0, 3 and 6 months) to evaluate their storage stability. The PV was determined through iodometric titration, following an AOAC method with slight modifications. Initially, the encapsulated oil was extracted with cold hexane for 24 h. Then, 0.20 ± 0.01 g of the extracted oil was mixed with 3 mL of acetic acid:chloroform (3:2% v/v) and stirred vigorously until it was completely dissolved. Next, 0.5 mL of saturated potassium iodide solution was added and the mixture was kept in the dark for 1 min. The reaction was stopped by adding 3 mL of distilled water, followed by 0.5 mL of starch solution (1%, w/v) as an indicator. Finally, the solution was titrated with 0.001 N Na₂SO₃ until the brown color disappeared. The PV was calculated in milliequivalents of oxygen per kilogram of oil using Equation (3). This experiment was conducted in triplicate for each sample.

PV = [(S − B) × N × 1000]/w

(3)

where S is the volume in mL of the Na₂SO₃ used by the sample, and B is the volume used by the blank. N is the normality of Na₂SO₃, and w represents the mass of oil expressed in grams.

A method used to measure oil properties is by determining the acid value (AV). The AOAC protocol for AV analysis in oils involves titration in ethanol using phenolphthalein as an indicator. To determine the AV, approximately 1 g of oil was dissolved in 8 mL of a solvent mixture with phenolphthalein (ethanol:diethyl ether = 1:1), and the mixture was shaken for 2 min. Next, it was titrated with 0.01 N KOH solution until the solution remained faintly pink after shaking for 30 s [27]. The acid value was then calculated using Equation (4), and it was expressed in milligrams of KOH required to neutralize the acidic constituents in 1 g of the sample.

Acid value = [V × 5.611]/Weight (g) of the sample

(4)

where V represents the volume in mL of the 0.01 N KOH solution and W represents the mass of the sample in grams.

2.13. Statistical Analysis

The study data are presented as the mean ± standard deviation (S.D.). Statistical analyses were conducted using one-way analysis of variance (ANOVA) and followed by Duncan’s post-hoc test. The significance level was set at α = 0.05.

3. Results and Discussion

3.1. Proximate Analysis of Chia Seeds and Fatty Acid Composition

Proximate analysis of chia seeds obtained from the Plant Breeding Center for Sustainable Agriculture, Faculty of Agriculture, Khon Kaen University, Thailand, showed a comparatively similar nutrient composition to commercially available chia previously reported by [28]. The KKU chia seeds had moisture content of 5.56 ± 0.08%, ash 4.83 ± 0.01%, lipid 31.29 ± 0.01%, crude fiber 20.67 ± 0.55% and protein 25.82 ± 0.25% respectively. Moreover, KKU chia seeds showed higher protein content compared to the commercial chia in the earlier study by Silva et al. [29]. Despite not being commonly cultivated as a protein source, chia seeds contain more protein than conventional crops such as corn, wheat, rice and oats [30]. Additionally, the fiber found in chia seeds induces satiety and retards digestion, resulting in a gradual increase in blood sugar levels and the more consistent release of insulin [3]. Both the soluble and insoluble dietary fiber present in chia seeds play a crucial role in digestive health and offer nutritional and physiological benefits to consumers, which are associated with lowering the risk of hypertension, coronary heart disease, obesity and diabetes [31].

The interest in chia seeds lies in their oil content, which is a rich source of polyunsaturated fatty acids (PUFAs). Chia oil contains both saturated fatty acids (SFA) and PUFAs, with α-linolenic acid being the most abundant and interesting PUFA present. The fatty acid profile of KKU chia oil predominantly consists of α-linolenic acid, as well as linoleic, oleic, palmitic and stearic acids. These findings are consistent with earlier studies [32]. GC chromatography and retention times reveal the fatty acid composition of KKU chia oil, with palmitic acid (C16:0) at 4.315 min, stearic acid (C18:0) at 6.128 min, oleic acid (C18:1) at 6.348 min, linoleic acid (C18:2) at 6.823 min and α-linolenic acid (C18:3) at 7.549 min. α-Linolenic acid is the major fatty acid present in chia oil, comprising 53.67 g/100 g oil. GC-MS analysis confirmed the fatty acid composition of KKU chia seeds, although the study revealed lower α-linolenic acid content compared to the previous study by Bodoira et al. [33], which reported up to 61.8 g/100 g oil.

The fatty acid composition of chia oil may differ depending on the variety and planting of chia seeds. Ayerza and Coates identified only four fatty acids, namely α-linolenic, linoleic, oleic and palmitic acids, while another study by Pintado et al. [34] found eight fatty acids, namely myristic, palmitic, stearic, oleic, linoleic, α-linolenic, arachidic and behenic acid. Chia oil has a higher percentage of polyunsaturated fatty acids (73.93%) than flaxseed oil (71.8%), mustard (31.5%), soybean (59.8%), rice bran (35.9%), corn (57.1%), sesame (45.7%), olive (10%) and sunflower (66%), with the exception of safflower (77.3%). However, safflower oil contains less than 1% ω-3. The n-6/n-3 fatty acid ratio of KKU chia oil (0.37) is similar to that found in a Chilean chia oil study (0.29) conducted by Marineli et al. [3]. The n-6/n-3 ratio in chia oil is notably lower than that in most vegetable oils, such as maize (76.57), rapeseed (2.26), soybean (6.68), sunflower (30.77) and olive (17.86) [35]. A low n-6/n-3 fatty acid ratio has been linked to a reduced risk of cardiovascular disease [36]. The MUFA/PUFA ratio of KKU chia oil (0.13) is higher than that of Chilean chia oil (0.08) due to its higher α-linolenic acid content.

3.2. Encapsulation Formulas and Chemical Properties

The formation of emulsions is a critical stage in the process of microencapsulating oils. The emulsions were prepared by using the ingredients of chia oil, which act as the core material, and a combination of WP and GA as the wall material. According to a previous study by Calvo et al. [8], the optimal encapsulation efficiency was achieved using a core material to wall material ratio of 1:5. For this study, six different wall materials were created using whey protein and gum arabic at varying ratios of 1:0, 1:1, 1:2, 2:1, 1:3 and 3:1 (w/w), as shown in Table 1. To ensure the complete dissolution of the wall materials, they were blended in double-distilled water before adding the chia oil. The stability and droplet size of the emulsions are crucial factors in determining the encapsulation efficiency and microcapsule morphology. Most studies on the microencapsulation of oils rich in unsaturated fatty acids have focused on using fish or flaxseed oils as the core material. However, there is currently growing interest in microencapsulating chia oil for use as a supplement in food and health products. One study by Ayerza et al. [7] examined the microencapsulation of chia oil using binary blends of WP concentrate with either GA or mesquite gum as the wall material.

In our study, six emulsion formulas were dried by SD or FD to obtain the microencapsulated powders. The results showed that solid yields from drying by SD and FD were 40.28–48.18 g and 60.86–68.06 g, accounting for 45.57–54.50% and 68.85–76.99%, respectively (

Table 2. Physicochemical characteristics of the microcapsules containing chia seed oil.

Formula	Yield (g)	Yield (%)	Moisture Content (wt%)	Water Activity (aw)	Color
					L*	a*	b*
W10S	43.86	49.62	3.13 ± 0.06 e	0.11 ± 0.04 a	92.24 ± 0.02 f	0.24 ± 0.01 f	26.96 ± 0.01 g
A11S	46.40	52.49	3.41 ± 0.05 bcd	0.15 ± 0.05 a	94.99 ± 0.06 c	−0.79 ± 0.02 i	23.59 ± 0.01 j
B12S	41.04	46.43	3.20 ± 0.07 de	0.12 ± 0.02 a	95.43 ± 0.06 b	−0.99 ± 0.01 j	22.86 ± 0.01 k
C21S	40.28	45.57	3.59 ± 0.09 ab	0.16 ± 0.02 a	93.68 ± 0.02 d	−0.42 ± 0.01 h	24.02 ± 0.01 i
D13S	45.18	51.11	3.40 ± 0.07 bgd	0.14 ± 0.01 a	96.45 ± 0.02 a	−1.61 ± 0.01 k	21.51 ± 0.01 l
E31S	48.18	54.50	3.20 ± 0.08 de	0.12 ± 0.02 a	93.23 ± 0.05 e	−0.07 ± 0.01 g	25.58 ± 0.01 h
W10F	60.86	68.85	3.36 ± 0.10 bcde	0.13 ± 0.01 a	86.10 ± 0.01 j	3.84 ± 0.01 b	38.37 ± 0.01 a
A11F	61.90	70.02	3.50 ± 0.08 abc	0.12 ± 0.04 a	85.73 ± 0.07 k	2.33 ± 0.01 c	30.39 ± 0.02 c
B12F	62.82	71.06	3.13 ± 0.09 e	0.14 ± 0.03 a	86.27 ± 0.01 i	1.26 ± 0.01 e	27.94 ± 0.01 e
C21F	68.06	76.99	3.70 ± 0.11 a	0.18 ± 0.06 a	87.37 ± 0.01 g	2.29 ± 0.01 d	28.73 ± 0.01 d
D13F	64.64	73.12	3.40 ± 0.08 bcd	0.13 ± 0.05 a	86.52 ± 0.01 h	1.27 ± 0.01 e	27.76 ± 0.01 f
E31F	67.20	76.02	3.31 ± 0.06 cde	0.13 ± 0.03 a	82.84 ± 0.05 l	3.97 ± 0.01 a	31.99 ± 0.02 b
WP					85.41 ± 0.01	3.89 ± 0.01	39.26 ± 0.02
GA					97.41 ± 0.19	−1.78 ± 0.00	20.84 ± 0.03

The small letters represent the significant level for the mean within the same column. If two means in each column share the same letter, they are not significantly different (p ≤ 0.05). W10S, A11S, B12S, C21S, D13S and E31S samples were dried with spray-drying process; W10F, A11F, B12F, C21F, D13F and E31F samples were dried with freeze-drying process. WP: whey protein, GA: gum arabic.

).

An analysis of the drying processes was performed through the examination of the recovery solid yields. The results indicated that the solid yields of the spray-dried (SD) samples were noticeably lower than those of the freeze-dried (FD) samples. The lower solid yields of the SD samples could be attributed to the retention of a substantial amount of solid on the walls of the drying chamber during the drying process. This could be due to the relatively low temperature used in the SD process to maintain the chemical and functional properties of chia oil.

Micrographs of the chia oil microcapsules (Figure 1) were examined to compare the differences in the microcapsules achieved by the two different drying methods. The microcapsules obtained through the spray-drying (SD) method showed a fine powder with a spherical shape, continuous and homogeneous surface structure, low roughness and no visible pores or fractures. Similar morphologies have been reported in the literature for other SD materials. The particle size of the chia oil microcapsule powder in this study was less than 30 μm. In a previous study [1], a narrower range of droplet sizes was reported in emulsions containing chia oil and mixtures of whey protein and gum arabic at different concentrations as a wall material, with droplets ranging from 2.97 to 9.01 μm in diameter. On the other hand, microcapsules using soy protein and maltodextrin as wall materials exhibited a particle size range varying from 4 to 10 μm.

In this study, the samples obtained through the freeze-drying (FD) method had distinct shapes compared to the spray-drying method. The samples from the FD method had a flake-like structure with porous surfaces. Similar flake-like but porous structures have been reported in the literature for similar materials [11], and this structure has also been observed in earlier studies on materials prepared by FD [37]. We also noticed that the microcapsules were prone to easy fracturing, indicating that proper care must be taken to avoid capsule rupture and prevent moisture from entering and affecting the sample’s dryness. The absence of porosity in the microcapsules is important for preserving their oxidation resistance.

Table 2 summarizes the moisture content, water activity and color as determined by the two different drying procedures. In the present work, no significant differences in moisture content and water activity were observed between SD and FD microcapsules or among microcapsule formulas. The water content of the microcapsule powders obtained was around 3.1 to 3.7% (w/w). The FD powders had higher average moisture content than SD powders. This could be a result of the relatively long drying period and greater pore formation during the FD process, allowing moisture to be more easily absorbed into the pores of the microcapsules. However, both microcapsule formulas without GA had a negligible amount of moisture. It is possible that the water-binding ability of GA in each formulation plays a role in retaining moisture in the dried powder. The main color differences in the chia microcapsule powder were due to the way in which the microcapsules were dried, the ratio of wall material in each formula, the particle size and the shape of the powder. All encapsulation samples exhibited high L* values (having a white and luminous shade). The L* values of SD microcapsules were brighter than those of FD microcapsules and their brightness was similar to the GA color, but the L* values of FD microcapsules were similar to the WP color. In addition, it has been recognized that the particle size and shape of SD microcapsules are constantly spherical. Therefore, the more scattered the light of the particle is, the brighter the sample will look. Conversely, the porous sheet shape of the particles obtained from FD caused the reflected light to reflect unevenly. This made the light appear darker. Gum arabic used as one of the wall materials had a significant influence on the a* and b* values of SD. The lower ratio of GA in the microcapsules caused higher a* and b* values, revealing a higher redness intensity and a yellow appearance of the SD microcapsules. In our study, the formula with the highest ratio of GA was D13S, which had the least redness (more greenness) and yellowness (more blueness). The a* and b* values of D13S were similar to those of GA. However, the ratio of GA had no significant impact on the a* and b* values of FD microcapsules. In conclusion, both SD and FD could be used to produce chia oil microencapsulation powders with relatively low moisture content. However, the SD process may be more suitable for chia oil microencapsulation as it yields lower moisture content, which may contribute to a longer shelf-life.

3.3. Effect of Encapsulation Storage on Chemical Properties

Encapsulation efficiency (EE) is a crucial factor in determining the degree of oil protection in microencapsulation, which is influenced by several factors, such as the material to be encapsulated, wall material composition, particle size and drying conditions. Achieving high %EE, low surface oil and increased core stability is essential for successful microencapsulation. The microcapsules obtained through the SD and FD processes were stored in laminated bags in a vacuumed and dark environment for 6 months to examine the effect of time on the surface oil (SO) and %EE, as shown in

Table 3. Physicochemical characteristics of the microcapsules containing chia oil stored in vacuum-sealed laminated bags with no light.

Formula	0 Month		3 Months		6 Months
	SO (g/100 g)	%EE	SO (g/100 g)	%EE	SO (g/100 g)	%EE
W10S	2.25 ± 0.05 g	62.10 ± 0.32 b	2.26 ± 0.06 h	58.25 ± 0.43 c	2.75 ± 0.05 g	50.92 ± 0.72 c
A11S	2.22 ± 0.08 g	65.81 ± 0.49 a	2.22 ± 0.07 hi	60.70 ± 0.46 b	2.63 ± 0.05 ghi	52.24 ± 0.41 c
B12S	1.92 ± 0.06 h	57.07 ± 0.56 c	2.07 ± 0.06 i	56.26 ± 0.60 d	2.45 ± 0.06 i	45.91 ± 0.67 f
C21S	2.03 ± 0.04 h	65.51 ± 0.24 a	2.46 ± 0.06 g	62.80 ± 0.47 a	2.68 ± 0.07 gh	55.43 ± 0.70 ab
D13S	1.98 ± 0.03 h	61.95 ± 0.28 b	2.96 ± 0.06 f	58.41 ± 0.37 c	2.76 ± 0.06 g	54.63 ± 0.45 b
E31S	2.25 ± 0.04 g	65.06 ± 0.24 a	2.26 ± 0.06 h	59.37 ± 0.61 c	2.54 ± 0.08 hi	56.63 ± 0.60 a
W10F	3.53 ± 0.02 f	46.30 ± 0.26 g	3.64 ± 0.07 e	43.74 ± 0.55 h	4.04 ± 0.06 f	38.00 ± 0.43 h
A11F	6.22 ± 0.05 b	56.60 ± 0.17 cd	6.66 ± 0.08 b	54.56 ± 0.25 e	7.63 ± 0.05 a	47.40 ± 0.19 e
B12F	6.06 ± 0.04 c	56.69 ± 0.13 cd	6.27 ± 0.06 c	52.99 ± 0.21 f	6.87 ± 0.06 c	48.87 ± 0.21 d
C21F	6.61 ± 0.03 a	55.42 ± 0.10 de	7.02 ± 0.05 a	54.65 ± 0.15 e	7.23 ± 0.06 b	51.92 ± 0.18 c
D13F	5.22 ± 0.04 d	54.22 ± 0.16 e	5.35 ± 0.03 d	51.33 ± 0.18 g	5.88 ± 0.09 e	46.70 ± 0.42 ef
E31F	4.27 ± 0.05 e	51.07 ± 0.28 f	6.53 ± 0.04 b	50.54 ± 0.14 g	6.32 ± 0.08 d	39.73 ± 0.46 g

The small letters represent the significant level for the mean within the same column. If two means in each column share the same letter, they are not significantly different (p ≤ 0.05). SO: surface oil, EE: encapsulation efficiency.

. The results indicate that the whey-protein-based wall materials were effective in retaining oil (at an oil:wall material ratio of 1:5) during the freezing, sublimation and spray-drying processes, as the surface oil values were similar to those of the initially added oil (0.166 g/g solids).

In this study, we observed that the lipophilic core material was embedded within the solid wall matrix as microparticles or microdroplets. Oil release increased with the storage time, and higher surface oils had a significant impact on the %EE. We found that the %EE values were higher in emulsions with a variation of GA, and, in formulations with lower GA ratios, the surface oil was significantly lower and %EE was significantly higher. The %EE of SD microcapsules was significantly higher than that of FD microcapsules, and the %EE values of SD microcapsules were significantly different among wall component proportions. The surface oil of SD microcapsules was significantly lower, and %EE was significantly higher in microcapsules containing lower GA than those containing higher GA. We also found that the ratio of WP to GA did not affect the SO and %EE, and that the particle size variations of the microcapsules did not affect the SO and %EE during storage. Our findings are consistent with those reported by Ayerza et al. [38], but differ from those reported by [35], who suggested that %EE could be improved by having narrowly distributed powder particles with an optimal size, which was <38 µm for non-volatiles (fish oil) and between 38 and 63 µm for volatiles (D-limonene). In the present work, we found that FD microcapsules stored at 0, 3 and 6 months had %EE values of 46.30–56.69%, 43.74–54.65%, and 38.00–51.92%, and SD microcapsules had %EE values of 57.07–65.81%, 56.26–62.80% and 45.91–56.63% respectively. Therefore, the C21S and E31S formulas were the most suitable emulsion formulas for chia oil microcapsule preparation due to having the highest %EE. Our results are consistent with those reported by González et al. [1], who found that %EE values ranged from 52.5 to 60.2 in the SD process and from 59.6 to 65.5 in the FD process. Similarly, González reported that %EE was over 70% for all microcapsules, and that both binary biopolymer blends produced parent and reconstituted emulsions that were stable against droplet coalescence. Previous research has suggested that an increase in the total solid concentration can result in higher viscosity and the formation of smaller droplets in the emulsion, which reduces the circulation of oil inside the particles, preventing its migration to the surface and thereby improving oil encapsulation [14]. It has also been reported that a combination of proteins and polysaccharides can increase emulsion stability and lead to higher encapsulation efficiency, depending on the nature of the biopolymers and the degree of complexity.

All microcapsules were stored in laminated bags in dark and vacuumed conditions for 3 and 6 months to examine the impact of time on the oxidative stability of the encapsulated oil by determining the fatty acid composition, AV and PV. The results revealed that microcapsules contained stearic, palmitic, oleic, linoleic and α-linolenic acid. α-Linolenic acid was the most detected fatty acid, found at more than 50%, followed by linoleic acid at approximately 20%, with the lowest being stearic acid at approximately less than 5%. The findings are shown in

Table 4. Chemical characteristics of chai oil containing microcapsules stored in the laminated bags under vacuum for 6 months.

Fatty Acid	Month	Formula
		W10S	A11S	B12S	C21S	D13S	E31S
C16:0	0	8.24 ± 0.77 ij	10.41 ± 0.37 b–g	9.11 ± 0.37 f–i	8.56 ± 0.35 h–j	8.87 ± 0.31 g–j	9.09 ± 0.24 g–i
	3	11.23 ± 0.30 a–d	9.69 ± 1.20 d–i	10.36 ± 0.46 b–g	9.25 ± 1.09 e–i	10.48 ± 1.56 b–g	11.76 ± 0.75 ab
	6	11.57 ± 0.64 a–c	9.63 ± 0.62 d–i	8.92 ± 1.57 g–j	9.57 ± 0.66 e–i	9.96 ± 2.30 c–h	10.86 ± 1.64 b–e
C18:0	0	4.86 ± 0.14 a	3.47 ± 0.09 e–k	3.56 ± 0.04 e–k	3.62 ± 0.17 d–k	4.66 ± 0.52 a–c	3.54 ± 0.06 e–k
	3	3.73 ± 1.37 c–k	4.10 ± 0.57 a–h	4.37 ± 0.58 a–e	4.53 ± 0.06 a–d	3.69 ± 0.97 d–k	3.39 ± 1.30 f–k
	6	3.72 ± 0.76 c–k	4.02 ± 0.69 a–i	4.75 ± 0.85 ab	4.37 ± 0.33 a–e	3.86 ± 0.96 b–j	2.87 ± 1.18 k
C18:1	0	9.05 ± 0.08 c–h	7.67 ± 0.43 i	8.87 ± 0.04 d–h	8.82 ± 0.23 d–i	8.04 ± 0.26 hi	9.13 ± 0.41 c–h
	3	9.66 ± 1.24 b–e	9.29 ± 0.47 b–g	9.24 ± 0.41 c–g	9.54 ± 0.59 b–f	8.87 ± 0.24 d–h	9.05 ± 0.16 c–h
	6	10.40 ± 1.38 ab	9.93 ± 0.66 a–d	11.02 ± 2.02 a	9.04 ± 0.15 c–h	9.17 ± 0.93 c–h	9.32 ± 2.04 b–g
C18:2	0	20.21 ± 0.35 b–g	20.10 ± 0.06 b–h	20.36 ± 0.16 b–e	20.79 ± 0.07 a–c	20.15 ± 0.06 b–g	20.28 ± 0.17 b–f
	3	19.57 ± 0.82 c–i	20.67 ± 1.61 a–c	19.10 ± 0.19 d–i	21.03 ± 1.73 ab	20.65 ± 1.68 a–c	20.16 ± 1.27 b–g
	6	18.66 ± 0.09 ij	19.02 ± 1.12 e–i	17.52 ± 1.20 j	21.90 ± 2.00 a	18.87 ± 0.96 g–j	19.15 ± 0.67 d–i
C18:3	0	57.64 ± 0.49 d–i	58.35 ± 0.03 b–f	58.10 ± 0.28 d–g	58.21 ± 0.11 c–g	58.27 ± 0.11 b–f	57.96 ± 0.06 d–h
	3	55.80 ± 1.64 j–l	56.25 ± 0.21 h–l	56.93 ± 0.54 e–j	55.64 ± 1.76 j–l	56.31 ± 0.68 h–l	55.64 ± 1.00 j–l
	6	55.65 ± 1.34 j–l	57.40 ± 1.05 d–j	57.79 ± 0.54 d–i	55.12 ± 0.91 kl	58.13 ± 1.40 d–g	57.80 ± 2.12 d–i
MUFA/PUFA	0	0.12	0.10	0.11	0.11	0.10	0.12
	3	0.13	0.12	0.12	0.12	0.12	0.12
	6	0.14	0.13	0.15	0.12	0.12	0.12
Fatty Acid	Month	Formula
		W10F	A11F	B12F	C21F	D13F	E31F
C16:0	0	9.40 ± 0.66 e–i	7.40 ± 0.12 j	10.36 ± 1.45 b–g	9.06 ± 0.51 g–i	9.10 ± 0.52 g–i	8.65 ± 0.16 h–j
	3	11.78 ± 0.94 ab	9.42 ± 0.99 e–i	9.29 ± 1.22 e–i	11.72 ± 1.02 ab	9.17 ± 1.55 f–i	10.74 ± 0.55 b–f
	6	11.36 ± 1.12 a–c	9.22 ± 1.25 f–i	9.19 ± 1.20 f–i	12.72 ± 0.88 a	9.55 ± 0.69 e–i	10.32 ± 1.42 b–g
C18:0	0	4.27 ± 0.25 a–f	4.08 ± 0.21 a–h	3.20 ± 0.11 h–k	4.33 ± 0.27 a–f	3.58 ± 0.05 e–k	4.00 ± 0.38 a–j
	3	3.58 ± 0.92 e–k	3.85 ± 0.55 b–j	3.53 ± 1.29 e–k	3.93 ± 0.84 a–j	4.65 ± 0.13 a–c	4.14 ± 0.49 a–h
	6	3.28 ± 0.85 g–k	3.06 ± 0.98 jk	3.31 ± 0.80 g–k	3.53 ± 0.66 e–k	4.19 ± 0.12 a–g	3.12 ± 0.71 i–k
C18:1	0	8.38 ± 0.04 g–i	8.44 ± 0.05 g–i	9.11 ± 0.08 c–h	8.84 ± 0.37 d–h	8.49 ± 0.01 f–i	8.38 ± 0.20 g–i
	3	9.00 ± 0.19 c–h	9.02 ± 0.18 c–h	8.46 ± 1.18 f–i	10.05 ± 1.24 a–c	9.04 ± 0.57 c–h	9.59 ± 0.61 b–f
	6	9.68 ± 0.83 b–d	8.25 ± 0.15 g–i	8.52 ± 0.29 e–i	10.05 ± 1.39 a–c	9.39 ± 0.79 b–g	9.14 ± 0.66 c–h
C18:2	0	20.25 ± 0.06 b–g	20.13 ± 0.21 b–g	20.05 ± 0.17 b–h	19.86 ± 0.07 b–i	20.40 ± 0.04 b–e	20.22 ± 0.03 b–g
	3	19.89 ± 0.77 b–i	19.93 ± 0.72 b–i	20.07 ± 1.26 b–h	21.04 ± 2.10 ab	19.74 ± 1.43 b–i	19.47 ± 0.82 c–i
	6	18.92 ± 0.18 f–i	19.48 ± 0.76 c–i	18.72 ± 0.71 h–j	19.13 ± 0.46 d–i	20.42 ± 1.75 b–d	18.91 ± 0.44 f–i
C18:3	0	57.70 ± 0.39 d–i	59.95 ± 0.08 a–c	57.28 ± 1.15 d–j	57.91 ± 1.07 d–h	58.43 ± 0.52 b–f	58.76 ± 0.77 a–d
	3	55.75 ± 0.69 j–l	57.78 ± 0.65 d–i	58.65 ± 1.67 a–e	53.26 ± 2.73 m	57.40 ± 1.04 d–j	56.06 ± 0.96 i–l
	6	56.75 ± 0.93 f–k	60.00 ± 1.35 ab	60.25 ± 1.17 a	54.58 ± 2.45 lm	56.45 ± 1.40 g–k	58.52 ± 1.26 a–f
MUFA/PUFA	0	0.11	0.11	0.12	0.11	0.11	0.11
	3	0.12	0.12	0.11	0.14	0.12	0.13
	6	0.13	0.10	0.11	0.14	0.12	0.12

The small letters represent the significant level for the mean within the same row, or within the same column. Any two means within the same row or the same column followed by the same letter are not significantly different (p ≤ 0.05). Means followed by different letters within the same row or the same column are significantly different based on LSD at p ≤ 0.05. W10S, A11S, B12S, C21S, D13S and E31S samples were dried with spray-drying process; W10F, A11F, B12F, C21F, D13F and E31F samples were dried with freeze-drying process.

. These results agree with those reported in previous studies [32,38,39]. However, in a previous study, the extracted chia oil from the capsule had a similar MUFA/PUFA ratio to chia oil. Therefore, encapsulation can protect against the oxidation of unsaturated fatty acids. According to a study by Sardenne et al. [40], the MUFA/PUFA ratio of Chilean chia oil (0.08) was notably lower than that of KKU chia oil (0.13) and lower than that of most other vegetable oils. Incorporating chia oil into one’s daily diet may offer several health benefits, as vegetable oils that contain high levels of PUFAs have been shown to be beneficial [12]. Moreover, it was found that the microcapsules W10S, B12S, D13S, A11F, B12F and E31F contained the highest ω-3. However, after 3 and 6 months of storage, the ω-3 was still higher in the A11F and B12F formulas. Therefore, the best combination is to prepare emulsion formulas A11F or B12F, followed by drying with the FD method; this results in better oil quality retention with the highest ω-3 content. At 0 months, W10 microcapsules formulated without GA and dried by the SD method were found to have higher amounts of ω-3 when compared to the FD method. However, throughout 3 and 6 months of storage, the ω-3 of W10S substantially decreased, but this was not observed in W10F. W10F microcapsules with the FD method had constant ω-3 content throughout the 3- and 6-month storage period, indicating that the FD method has a lesser effect on fatty acid oxidation compared to the SD method of drying. In this case, the formula containing GA (A11F, B12F and E31F) with FD had ω-3 content that was relatively high and steady during storage compared to SD. Therefore, it is beneficial to prepare emulsion formulas A11, B12 or E31 to preserve the unsaturated fatty acid content. Chia oil contains three types of fatty acids: SFA, MUFA and PUFA. In this study, chia oil had a ratio of SFA:MUFA:PUFA at 1.59:1:7.34, which was different from the World Health Organization recommendation of a fatty acid ratio of 1:1.5:1 for vegetable oil. However, the body can build SFA itself and it should be obtained through normal, adequate daily intake, while the body cannot generate some PUFAs, including linoleic acid (ω-6) and α-linolenic acid (ω-3). These should be consumed every day to prevent deficiencies in essential fatty acids. Chia oil is among the highest ω-6 and ω-3 constituents. Moreover, MUFA, a good fatty acid, is also found in chia oil (although in small amounts of approximately 10%) [41]. The body needs MUFA because it helps in reducing bad cholesterol (LDL) and increasing good cholesterol (HDL).

The acid value (AV) is an important characteristic used to describe fats and oils. It refers to the quantity of KOH in mg required to counteract the organic acids found in one gram of oil, and it reflects the concentration of free fatty acids (FFA) present in the oil.

Table 5. Acid values and peroxide values of the microcapsules containing chia oil at different storage times from the spray-drying and freeze-drying methods.

Formula	Acid Value (mg KOH)			Peroxide Value (Meq Peroxide/kg Oil)
	0	3	6	0	3	6
W10S	2.46 ± 0.30 b	3.13 ± 0.22 ab	3.72 ± 0.20 ab	1.33 ± 0.28 b	4.83 ± 0.76 ab	20.33 ± 0.57 abc
A11S	2.53 ± 0.42 b	2.90 ± 0.06 abcd	3.65 ± 0.25 ab	1.17 ± 0.28 b	5.50 ± 0.50 ab	20.83 ± 0.76 ab
B12S	2.37 ± 0.28 bc	2.92 ± 0.13 abc	3.78 ± 0.22 a	1.17 ± 0.28 b	5.50 ± 0.86 ab	20.83 ± 0.76 ab
C21S	2.70 ± 0.43 ab	3.17 ± 0.18 a	3.68 ± 0.17 ab	2.67 ± 0.76 a	4.83 ± 1.04 ab	21.33 ± 1.04 a
D13S	3.36 ± 0.46 a	3.13 ± 0.20 ab	3.49 ± 0.11 ab	3.33 ± 0.28 a	6.00 ± 0.00 a	22.00 ± 0.86 a
E31S	3.10 ± 0.21 ab	3.24 ± 0.08 a	3.64 ± 0.24 ab	3.33 ± 0.28 a	5.33 ± 1.04 ab	20.50 ± 0.50 abc
W10F	1.56 ± 0.11 cd	2.50 ± 0.23 cd	3.03 ± 0.15 cd	1.17 ± 0.28 b	4.50 ± 0.50 ab	17.50 ± 0.86 d
A11F	1.31 ± 0.17 d	2.65 ± 0.10 cd	2.93 ± 0.12 d	1.17 ± 0.28 b	3.83 ± 0.76 b	18.17 ± 1.04 cd
B12F	1.41 ± 0.20 d	2.47 ± 0.22 d	3.23 ± 0.09 bcd	1.00 ± 0.00 b	4.33 ± 0.76 ab	18.17 ± 0.76 cd
C21F	1.42 ± 0.17 d	2.45 ± 0.17 d	3.05 ± 0.18 cd	1.17 ± 0.28 b	4.50 ± 0.50 ab	18.50 ± 1.00 bcd
D13F	1.33 ± 0.06 d	2.71 ± 0.05 bcd	2.97 ± 0.11 d	1.17 ± 0.28 b	4.67 ± 0.57 ab	17.83 ± 1.60 d
E31F	1.34 ± 0.09 d	2.48 ± 0.12 d	3.03 ± 0.24 cd	1.50 ± 0.00 b	4.17 ± 0.28 ab	18.17 ± 0.57 cd

The small letters represent the significant level for the mean within the same column. If two means in each column share the same letter, they are not significantly different (p ≤ 0.05). W10S, A11S, B12S, C21S, D13S and E31S samples were dried with spray-drying process; W10F, A11F, B12F, C21F, D13F and E31F samples were dried with freeze-drying process.

shows that the microcapsules of the SD method gave higher acid values than those of the FD method. This may be due to the use of heat during the sample drying process. From this result, therefore, FD would be a better drying option as it reduces the AV, although it requires a longer drying time. However, every storage sample had an AV no more than the Codex limit of 40 mg KOH/g oil. Table 5 shows that the peroxide values (meq peroxide/kg oil) of the microcapsules containing chia oil from the SD and FD methods gradually increased with storage time. Of all formulas in each storage period, the PVs of powders with the SD method were higher than those with the FD method, except formula A11 at 0 months. The formulas with the highest PVs were D13S, E31S and C21S at 0 months. At 3 months, the highest PVs was found for D13S but it was not statistically different from other formulas. At 6 months, the highest PVs were noted for D13S and C21S, but the values were not statistically different from those of other formulas under drying with the SD method. Therefore, the microcapsules should be prepared and then dried by FD, since this method has low oxidation and PV values. The formula D13S should not be chosen because it had a high PV during 0–6 months of storage. The increase in the PV might be due to the ratio of WP to GA being 1:3, which did not show the ability to encapsulate chia oil efficiently. Although possessing good powder properties when dried by SD, giving a round shape, the chia oil was oxidized by heat while drying. As a result, the longer storage of encapsulated powders will cause more oxidation and a higher PV, i.e., storing the oil in powder for a long time will lead to more peroxide.

4. Conclusions

This study focuses on the microencapsulation of chia oil, which is an essential source of omega-3 PUFAs, particularly α-linolenic acid. The microencapsulation process described here involves the production of omega-3-rich powders that could be used as ingredients in enriched staple foods. The study compared the physicochemical properties of the microcapsules produced by the SD and FD methods and evaluated different wall materials, such as WP and GA, at different ratios to identify a stable encapsulated powder that can prevent the oxidation degradation of chia oil.

The investigation assessed the physicochemical and morphological properties, oxidative stability and storage life of the oil in all twelve microencapsulation wall materials. Differences were observed, and the quality of the encapsulation was evaluated through total oil and %EE analysis. %EE was found to be dependent on the ratio of wall material, showing the highest %EE at a WP to GA ratio of 2:1 and drying by SD methods over a storage period of 6 months.

During the storage period, the %EE values gradually decreased, implying a reduction in the storage life of the microencapsulated chia oil powder. However, encapsulated powders containing approximately 20% of chia oil and stored for six months showed both AV and PV values below the Codex limit, indicating the good quality of the encapsulated oil. Nevertheless, the study only evaluated the stability of the encapsulated powder for six months, which may not be enough to determine the long-term storage stability of the product. This study also demonstrated that microencapsulation provided effective protection from lipid oxidation by the wall matrix during the six-month storage period. In conclusion, the use of microencapsulation can enable chia oil to be applied in various fields, such as food production, nutraceuticals and cosmetics. The encapsulated chia oil has the potential to be incorporated into a variety of products, such as baking mixes, protein bars, skin creams and more.