Extraction and Microencapsulation of Phytochemical Compounds from Mango Peel (Mangifera indica L.) var. “Kent” and Assessment of Bioaccessibility through In Vitro Digestion

Abstract

The peel from mango (Mangifera indica L.) var. “Kent” is a good source of bioactive compounds (BC). BC are sensitive to oxygen, temperature, humidity, light, and gastrointestinal digestion, which change their biological function and health benefits. This study was aimed at the extraction of the bioactive compounds present in the peel from mango var. “Kent” and their microencapsulation using spray drying (SD) and spout-fluid bed drying (SFB). The bioaccessibility of BC was also evaluated. Two consecutive extractions of 90 min at 30 °C and 80% v/v ethanol were used. The microcapsules produced via SD and SFB presented high retention and encapsulation percentages of the bioactive compounds; nevertheless, SFB showed better protection during in vitro gastrointestinal digestion. The non-encapsulated extract showed a decrease (p ≤ 0.05) of BC at the end of in vitro gastrointestinal digestion. The results show that these microcapsules might be used in the food industry as an ingredient to produce functional foods and, thereby, to obtain the health benefits that the bioactive compounds provide.

1. Introduction

Mango (Mangifera indica L.) is a very important tropical fruit worldwide, primarily for its aroma, flavor, and attractive color. In Mexico, mango ranks among the top three most consumed fruits, followed by bananas and apples [1,2].

The main mango varieties are Manila, Ataulfo, Haden, Kent, Keiit, and Tommy Atkins. The mangoes known as “Manila” have medium-sized fruit and are orange and yellow in color, whereas the mangoes known as “petacones” (Haden, Kent, Keiit, and Tommy Atkins) have medium- to large-sized fruit, with rounded to ovoid shapes and a green-red-purple color [3]. Among the different cultivars of “petacón” mangoes, the “Kent” variety is one of the varieties most widely accepted by consumers, due to its excellent sensory attributes and appealing color. This variety has an oval shape with a rounded base, measuring 11 to 13 cm in length and weighing between 480 and 650 g. It ripens in red and yellow hues and has low fiber content, a sweet taste, and an average yield ranging between 100 and 300 kg per tree. The sweetness and juiciness of this mango variety make it suitable for juice extraction and for frozen mango products such as frozen pulp or chunks [1].

During mango processing, the peel is a waste product that accounts for 15 to 20% of the fruit’s weight. Mango peel can provide dietary fiber, energy, protein, carbohydrates, and fats. Additionally, it is an excellent source of phytochemical compounds like phenolic compounds, flavonoids, anthocyanins, and carotenoids. These phytochemicals act as antioxidants in the organism, thereby offering beneficial effects on health, including protection against chronic diseases, anti-inflammatory functions, and enhancement of immune response [2,4]. However, several of these phytochemical compounds are sensitive to conditions during food processing, environmental factors (temperature, light, humidity, and oxygen), and conditions in the gastrointestinal tract (pH, temperature, and enzymes), which could potentially impact the stability of these phytochemical compounds, thereby changing their biological activities and their possible beneficial effects [5].

The positive properties of phytochemicals can be assessed via bioaccessibility, which refers to the amount or portion of a phytochemical compound that is free to be absorbed in the small intestine. Phytochemical compounds will promote positive biological activities when they are absorbed at the end of gastrointestinal digestion [5].

In order for phytochemical compounds to have good biological effects on the organism, protective mechanisms are needed to preserve the active form of the compound until the moment of intake, digestion, and absorption. The microencapsulation process has been used to shield and prevent the degradation of phytochemical compounds [6]. Microencapsulation provides a barrier to shield phytochemical compounds from adverse conditions such as oxygen, light, heat, moisture, the digestive process, and the host’s immune system. Furthermore, microencapsulation enables the controlled release of the encapsulated product. Spray drying (SD) is a popular method used for the microencapsulation process. Additionally, there is a lower-cost alternative known as spout-fluid bed drying (SFB). Both drying techniques have been used to microencapsulate phytochemical compounds extracted from agro-industrial waste. The major advantage of the microencapsulation process is that it can be used with heat-resistant and heat-sensitive materials [6].

Various studies have been conducted to quantify phytochemical compounds and their antioxidant capacity in the peel of mangoes of different varieties [7,8,9,10,11,12]. These works have confirmed that mango peel has a high level of phytochemicals and high antioxidant capacity. No toxicity has been reported for mango peel [13]; thus, it can be used with confidence to extract phytochemical compounds that could serve as ingredients to produce functional foods, nutraceuticals, or functional beverages that can confer health benefits.

There are few studies on the peel of “Kent” variety mangoes [14,15,16] and, currently, no works have described the microencapsulation of phytochemical compounds of “Kent” variety mango peel by SD and SFB. Additionally, the in vitro bioaccessibility of microencapsulated peel phytochemical compounds from “Kent” variety mangoes has also not been analyzed.

The aim of this work was to extract and encapsulate the phytochemical compounds present in the peel of “Kent” variety mangoes (Mangifera indica L.) by SD and SFB. Also, the bioaccessibility of the resulting microencapsulated phytochemical was evaluated through in vitro gastrointestinal digestion.

The study’s hypothesis is the following: that microencapsulation will protect these bioactive compounds and they will be gradually released to be absorbed in the intestine.

2. Materials and Methods

2.1. Reagents and Solvents

The reagents and solvents used in the study were acquired from JT Baker (Baker-Mallinckrodt, Mexico City, México). Pepsin from porcine gastric mucosa (EC 3.4.23.1), α-amylase (EC 3.2.1.1), pancreatin from porcine pancreas (EC 232.468.9), and porcine bile (EC 232.369.0) were purchased from Sigma-Aldrich Chemical Co. (St. Louis, MO, USA). Gum arabic was acquired from Agricurama S.P.R. de R.L. de C.V. (Mexico City, Mexico).

2.2. Mango Samples

“Kent” variety mangoes were obtained from the “Central de Abastos”, Ecatepec de Morelos, State of Mexico. These mangoes corresponded to the 2022 harvest. The fruits were acquired at a stage of consumption constituting ripeness, without apparent mechanical damage, pathogens, or insect-induced damage.

2.3. Sample Conditioning

“Kent” variety mangoes were cleaned with sodium hypochlorite solution (1.5 % v/v) and distilled water. Then, the mangoes were dried in ambient air (20 ± 2 °C). Subsequently, the pulp, peel, and seed were removed by hand and the percentage of each fraction was calculated. Next, the peel was dried at 45 °C for 24 h in an oven (Blue Lindberg V0914A, Thermo Scientific, Waltham, MA, USA). The dried peel was ground in a food processor (Magic Bullet Deluxe, BX1573-23, Mexico City, Mexico) and sieved through a 60-mesh sieve (250 μm) to standardize the particle size.

2.4. Extraction of Phytochemicals

At first, the extraction of polyphenols was performed following the methodology of Martínez-Ramos [17]: 1.0 g of fresh or powdered mango peel, placed in 20 mL of ethanol (80%), was stirred constantly (1000 rpm) for 90 min at room temperature (20 ± 2 °C). The extract obtained from the fresh or powdered mango peel was analyzed to establish the contents of phenolic compounds (PC), flavonoids (FC), mangiferin (MC), anthocyanins (AC), carotenoids (CC), and ascorbic acid (AA) and antioxidant capacity (ascertained with ABTS and DPPH).

2.5. Optimization of Extraction Conditions

The initial extraction conditions were optimized to extract at least 90% of the PC and FC. To optimize the extraction process, the number of consecutive extractions needed to obtain the highest concentrations of PC and FC was determined; subsequently, a Box–Behnken design was developed to determine the temperature, aqueous ethanol concentration, and extraction time for the maximum extraction of PC and FC.

2.5.1. Number of Consecutive Extractions

As already mentioned above, the optimization of the extraction process was performed by first determining the number of consecutive extractions sufficient to extract the highest concentrations of PC and FC, following the methodology of Martínez-Ramos et al. [17]. At the end of 90 min of extraction time, the extract was centrifuged at 1350× g for 10 min and the supernatant was decanted, then another 20 mL of 80% ethanol was added to the solids for the next extraction of 90 min, with constant agitation at 1000 rpm. This procedure was repeated until five consecutive extractions were obtained. Each extraction was centrifuged at 1350× g for 10 min, filtered (Whatman # 1, Maidstone, UK), and kept at 4 °C until analysis (after no more than 8 days). The extracts were analyzed for PC and FC contents.

2.5.2. Box–Behnken Design

Once the number of consecutive extractions for maximum PC and FC derivation was obtained, a Box–Behnken design was carried out. The design was based on 15 extractions with three replicates at the central point. The effects of temperature (30, 35, and 40 °C), aqueous ethanol concentration (50, 65, and 80%), and extraction time (30, 60, and 90 min) were analyzed. The Box–Behnken design was realized using Minitab 18.

2.6. Analytical Methods

2.6.1. Phenolic Compounds (PC)

PC were determined via the Folin–Ciocalteu method [18]. Absorbance was obtained at 765 nm (Jenway 6305, Long Branch, NJ, USA). The standard curve (0.1–200 mg/mL, n = 9, and R² = 0.9979) was constructed with gallic acid. The results were expressed as mg of gallic acid equivalent (GAE) per gram of dry weight (mg GAE/g_dw).

2.6.2. Flavonoid Content (FC)

FC was obtained following the methodology of Yin-Thoo et al. [19]. Absorbance was obtained at 510 nm (Jenway 6305, Long Branch, NJ, USA). A standard curve (0.1–1 mg/mL, n = 11, R² = 0.9908) was constructed with quercetin. The results were reported as mg of quercetin equivalent (QE) per gram of dry weight (mg QE/g_dw).

2.6.3. Mangiferin Content (MC)

MC was obtained following the methodology of Sumaya-Martínez et al. [15]. Absorbance was obtained at 410 nm (Jenway 6305, Long Branch, NJ, USA). A standard curve (0–200 µg mangiferin/mL DMSO (dimethyl sulfoxide), n = 8, and R² = 0.9861) was created with mangiferin. The results were expressed as mg of mangiferin equivalent (ME) per gram of dry weight (mg ME/g_dw).

2.6.4. Anthocyanin Content (AC)

For AC [20], absorbance was obtained at 510 and 700 nm (Jenway 6305, Long Branch, NJ, USA). The results were expressed as mg of cyanidin-3-glycoside per gram of dry weight (mg cya-3-glu/g_dw).

2.6.5. Carotenoid Content (CC)

CC was analyzed following the methodology of Singh-Sogi et al. [8]. The absorbance was obtained at 450 nm (Jenway 6305, Long Branch, NJ, USA). A standard curve (0–2.5 µg/mL hexane, n = 8, and R² = 0.9941) was constructed, using β-carotene as the standard. The results were expressed as mg of β-carotene per gram of dry weight (mg β-carotene/g_dw).

2.6.6. Ascorbic Acid Content (AA)

AA was determined using the colorimetric technique employed by Dürüst et al. [21]. Absorbance was obtained at 520 nm (Jenway 6305, Long Branch, NJ, USA). A standard curve (0.1–100 mg/L oxalic acid, 4%, n = 11, and R² = 0.9897) was created with ascorbic acid. The results were expressed as mg of ascorbic acid (AA) per gram of dry weight (mg AA/g_dw).

2.6.7. Antioxidant Capacity

Two free radicals were used to establish the antioxidant capacity: 2,2-azinobis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) and 2,2-diphenyl-1-picrylhydrazyl (DPPH).

For the ABTS method [22], absorbance was obtained at 734 nm (Jenway 6305, Long Branch, NJ, USA). The standard curve (0.1–1300 µmol, n = 8, and R² = 0.9926) was created with Trolox. The results were reported as µmol of Trolox equivalent per gram of dry weight (µmol TE/g_dw).

For the DPPH method [23], absorbance was obtained at 515 nm (Jenway 6305, Long Branch, NJ, USA). A standard curve (0–1300 µmol, n = 8, and R² = 0.9969) was created with Trolox. The results were expressed as µmol of Trolox equivalent per gram of dry weight (µmol TE/g_dw).

2.7. Microencapsulation

The extract was concentrated (4×) to eliminate ethanol and increase the PC to about 2 mg GAE/mL of extract to guarantee a high content of phenolic compounds in the microcapsules. To avoid precipitation of the wall material, all ethanol from the extract was removed using a rotavapor (BUCHI, R-300, Flawil, Switzerland) at 55 rpm, 40 °C, and 77 mBar.

A 30% w/w solution of gum arabic (GA) was prepared and left for 3 h in a vacuum oven (OV-12 Lab Companion, Seoul, Republic of Korea) at 40 °C. Next, the GA solution was mixed with the concentrated extract (final concentration: 10% w/w). This corresponded to a core:encapsulating agent ratio of 1:4.

2.7.1. Microencapsulation by Spray Drying (SD)

The microencapsulation process of SD was carried out in triplicate using a semi-pilot spray dryer (GEA-NIRO Atomizer, Mobile Minor, Copenhagen, Denmark), which was equipped with a two-fluid spraying nozzle and a cyclone separator to gather the powder. The air spraying pressure was 0.5 kg_f/cm². Microencapsulation was performed using the optimal conditions reported by Rigon and Zapata [24]: 170 °C inlet air temperature and 80 °C outlet air temperature. Afterward, the powders were stored at −18 °C.

2.7.2. Microencapsulation by Spout-Fluid Bed Dryer (SFB)

The microencapsulation process using an SFB with inert solids was carried out in triplicate with a spout-fluid bed dryer. This equipment had a cylindrical Perspex chamber with a perforated conical base and a short central tube submerged in the inert solids bed. The bed of inert solids consisted of a copolymer of fluor-ethylene-propylene pellets (FEP^®, DuPont Company, Wilmington, DE, USA). The equipment was operated with a nozzle spraying pressure of 0.5 kg_f/cm² and an inlet airflow velocity through the annular region that was equivalent to 0.7 U_mf (minimum fluidization velocity), with an inlet annular air temperature equal to the outlet drying temperature. The inlet airflow velocities through the spout were equivalent to 1.2 U_mst (minimum spouting velocity with a draft tube). Microencapsulation was performed using the following conditions: 160 °C of inlet temperature and 90 °C of outlet temperature. Afterward, the powders were stored at −18 °C.

2.8. Characterization of the Microcapsules

2.8.1. Total Phenolic Compounds on the Surface of the Microcapsules

This determination was made according to the method used by Robert et al. [25]. First, 0.8 g of microcapsules were mixed with 8 mL of a methanol:ethanol solution (1:1) and stirred with a vortex (MX-S, DLAB, Beijing, China) for 1 min. Subsequently, the mixture was filtered (Whatman^® #1). Finally, the filtrate was analyzed for PC.

2.8.2. Retention Efficiency (RE) and Encapsulation Efficiency (EE)

RE was determined by relating the PC of the reconstituted microcapsules to the PC in the initial solution to be dried (extract + wall material) per g of dry solids. RE was reported as a percentage, according to Equation (1):

$$\%\mathrm{R}\mathrm{E}=\left(\frac{\mathrm{P}\mathrm{C} \mathrm{i}\mathrm{n} \mathrm{r}\mathrm{e}\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{s}\mathrm{t}\mathrm{i}\mathrm{t}\mathrm{u}\mathrm{t}\mathrm{e}\mathrm{d} \mathrm{m}\mathrm{i}\mathrm{c}\mathrm{r}\mathrm{o}\mathrm{c}\mathrm{a}\mathrm{p}\mathrm{s}\mathrm{u}\mathrm{l}\mathrm{e}\mathrm{s}/{\mathrm{g}}_{\mathrm{d}\mathrm{w}}}{\mathrm{P}\mathrm{C} \mathrm{i}\mathrm{n} \mathrm{i}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l} \mathrm{s}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n} \mathrm{t}\mathrm{o} \mathrm{b}\mathrm{e} \mathrm{d}\mathrm{r}\mathrm{i}\mathrm{e}\mathrm{d}/{\mathrm{g}}_{\mathrm{d}\mathrm{w}}}\right)\times 100$$

(1)

EE was determined according to Equation (2):

$$\%\mathrm{E}\mathrm{E}=\left(\frac{\mathrm{P}\mathrm{C} \mathrm{i}\mathrm{n} \mathrm{r}\mathrm{e}\mathrm{c}\mathrm{o}\mathrm{n}\mathrm{s}\mathrm{t}\mathrm{i}\mathrm{t}\mathrm{u}\mathrm{t}\mathrm{e}\mathrm{d} \mathrm{m}\mathrm{i}\mathrm{c}\mathrm{r}\mathrm{o}\mathrm{c}\mathrm{a}\mathrm{p}\mathrm{s}\mathrm{u}\mathrm{l}\mathrm{e}\mathrm{s}/{\mathrm{g}}_{\mathrm{d}\mathrm{w}}-\mathrm{P}\mathrm{C} \mathrm{o}\mathrm{n} \mathrm{s}\mathrm{u}\mathrm{r}\mathrm{f}\mathrm{a}\mathrm{c}\mathrm{e} \mathrm{m}\mathrm{i}\mathrm{c}\mathrm{r}\mathrm{o}\mathrm{c}\mathrm{a}\mathrm{p}\mathrm{s}\mathrm{u}\mathrm{l}\mathrm{e}\mathrm{s}/{\mathrm{g}}_{\mathrm{d}\mathrm{w}}}{\mathrm{P}\mathrm{C} \mathrm{i}\mathrm{n} \mathrm{i}\mathrm{n}\mathrm{i}\mathrm{t}\mathrm{i}\mathrm{a}\mathrm{l} \mathrm{s}\mathrm{o}\mathrm{l}\mathrm{u}\mathrm{t}\mathrm{i}\mathrm{o}\mathrm{n} \mathrm{t}\mathrm{o} \mathrm{b}\mathrm{e} \mathrm{d}\mathrm{r}\mathrm{i}\mathrm{e}\mathrm{d}/{\mathrm{g}}_{\mathrm{d}\mathrm{w}}}\right)\times 100$$

(2)

2.8.3. Moisture Content and a_w (Water Activity)

To measure moisture content, a thermobalance (OHAUS MB23, Mexico City, Mexico) was used. a_w was obtained with an AquaLab Cx-3 (Decagon, Pullman, WA, USA).

2.8.4. Particle Size Distribution

Particle size and particle size distribution were analyzed with a Malvern laser diffraction particle size analyzer (IM 026 2006 series, Malvern, UK) using a 100 mm lens. The equivalent spherical diameter (D[4,3]) and Sauter diameter (D[3,2]) were analyzed using hexane (HONEYWELL, Mexico City, Mexico) as the dispersant fluid.

2.8.5. Morphology of Microcapsules

Morphology was acquired using a scanning electron microscope (SEM) (FEI Quanta FEG 250, Technical Cell, Kolkata, WB, India) at 2 kV. The microcapsule morphology was acquired at 1000× and 2000×.

2.9. In Vitro Digestion Procedure

The in vitro digestion procedure was based on the methodology described by Minekus et al. [5]. A control was also prepared (digestion without sample). The in vitro digestion process was carried out on the mango peel powder, concentrated extract, and microencapsulated extract using both drying techniques (SD and SFB).

The oral phase was simulated by mixing 5 g of the sample (mango peel powder, concentrated extract, or microcapsules) with 3.5 mL of salivary fluid (pH 7), 0.5 mL of α-amylase (75 U/mL), 0.025 mL of CaCl₂ (0.3 M), and 0.975 mL of distilled water. The mixture was stirred at 80 rpm at 37 °C for 2 min. At the end of 2 min, an aliquot was taken and kept on ice for 10 min. After the specified time elapsed, the aliquot was centrifuged at 1796× g for 10 min. Finally, the supernatant was analyzed for PC, FC, MC, AC, CC, AA, ABTS, and DPPH.

The gastric phase was simulated by mixing the bolus obtained from the oral phase with 7.5 mL of gastric fluid (pH 2), 1.6 mL of pepsin (2000 U/mL), 0.005 mL of CaCl₂ (0.3 M), 0.2 mL of HCl (1 M), and 0.695 mL of distilled water. The mixture was stirred at 80 rpm at 37 °C for 120 min. At the end of the 120 min, a 10 mL aliquot was taken and kept on ice for 10 min. After this time had elapsed, the aliquot was centrifuged at 1796× g for 10 min. Finally, the supernatant was analyzed for PC, FC, MC, AC, CC, AA, and antioxidant capacity (ABTS, DPPH).

The small intestine phase was simulated by mixing the remaining sample of gastric chyme with 5.5 mL of intestinal fluid (pH 7), 2.5 mL of pancreatin (100 U/mL), 1.25 mL of porcine bile extract (160 mM), 0.02 mL of CaCl₂ (0.3 M), 0.075 mL of NaOH (1 M), and 0.655 mL of distilled water. The mixture was stirred at 80 rpm at 37 °C for 120 min. At the end of the 120 min, the mixture was kept on ice for 10 min. After the time elapsed, the mixture was centrifuged at 1796× g for 10 min. Finally, the supernatant was analyzed for PC, FC, MC, AC, CC, AA, and antioxidant capacity (ABTS, DPPH).

The recovery (%R) and bioaccessibility (%B) percentages were evaluated according to Equations (3) and (4), respectively [26]:

$$\%R=\frac{A}{C}\times 100$$

(3)

$$\%B=\frac{B}{C}\times 100$$

(4)

where A is the amount of BC at the end of oral and gastric digestion. B is the amount of BC at the end of intestinal digestion. C is the amount of BC before in vitro gastrointestinal digestion.

2.10. Statistical Analysis

The results were evaluated with an ANOVA (one-way analysis of variance) and a comparison of means via the Tukey test (p ≤ 0.05) using the software Minitab 19 (Minitab, Inc., State College, PA, USA).

3. Results

3.1. Yield of “Kent” Mango

Each part of the fruit (pulp, seed, and peel) was weighed. The percentage corresponding to the peel was 16.87%, while the pulp was 70.41%, and the seed was 12.72%. An average moisture content of 74.84 ± 1.49% was obtained for the fresh peel, which coincides with the results of Marçal and Pintado [27]. Subsequently, the peel was dried, ground, and sieved to obtain the peel mango powder, which had a moisture content of 3.6 ± 0.06%. This value was lower than that reported elsewhere (5.8 ± 0.03%) by Marcillo-Parra et al. [4], who used freeze-drying as the drying method.

3.2. Bioactive Compounds (BC) and Antioxidant Capacity (AC) of Mango Peel

The characterization of the peel (fresh and dried powder) of the “Kent” mango variety is shown in

Table 1. Bioactive compounds and the antioxidant capacity of mango peel (fresh and dried powder).

Parameter	Fresh Mango Peel	Mango Peel Dried Powder
PC (mg GAE/gdw)	58.33 ± 2.97 a	57.57 ± 2.53 a
FC (mg QE/gdw)	11.35 ± 0.21 a	11.04 ± 0.16 a
MC (mg ME/gdw)	3.20 ± 0.23 a	3.01 ± 0.10 a
AC (mg cya-3-glu/gdw)	0.15 ± 0.01 a	0.14 ± 0.02 a
CC (mg β-carotene/gdw)	2.65 ± 0.08 a	2.38 ± 0.33 a
AA (mg AA/gdw)	12.66 ± 0.04 a	11.63 ± 0.12 b
Antioxidant capacity
ABTS (μmol TE/gdw)	144.86 ± 2.07 a,A	142.69 ± 0.31 a,A
DPPH (μmol TE/gdw)	185.07 ± 1.15 a,B	181.74 ± 0.22 b,B

Results are expressed as the mean ± standard deviation, n = 3. Different lowercase letters per row indicate a significant difference (p ≤ 0.05). Different capital letters per column indicate a significant difference (p ≤ 0.05) in antioxidant capacity.

. During the drying process, all the bioactive compounds and antioxidant capacity analyzed maintained the original concentrations (no significant difference; p ≤ 0.05) present in fresh mango peel, with the exception of AA, which lost 8.14% of its original content, showing a significant difference from the content in fresh mango peel.

The total phenolic compounds (PC) obtained from the fresh and dried mango peel (Table 1) are higher than those reported by other authors [4,15]. The same was true of the flavonoid content (FC). Marcillo-Parra et al. [4] reported an FC of 6 mg CE/g_dw in the “Kent” mango; however, the reference compounds are different. These authors based their results on catechin, while in this study, quercetin was used as the reference since Ajila et al. [7] and De Ancos et al. [10] reported quercetin as one of the main flavonoids present in mango.

Regarding the mangiferin content (MC) shown in Table 1, these results were higher than those reported by other authors [15,28]. The same was true for anthocyanins (AC), carotenoids (CC), and ascorbic acid (AA). The anthocyanin content of mango is markedly low, as anthocyanins are not responsible for the characteristic red, orange, and yellow tones of the fruit; instead, carotenoids are responsible for this color [10].

Differences between the bioactive compound contents shown in Table 1 for fresh and dried “Kent” mango peel with those reported by other authors may be attributed to factors such as the extraction process or fruit characteristics (ripeness and harvest season, among others) genetic base, agronomic practices, harvest stage, and location, among others [29].

Antioxidant capacity, measured with ABTS, was maintained during the drying process because there was no significant difference (p ≤ 0.05) between fresh and dried peel. These results were lower than those reported by other authors [2,4]. Antioxidant capacity, measured with DPPH, was higher than that obtained using ABTS. This may be due to the fact that mango peel has a higher level of lipophilic antioxidants since the DPPH technique primarily measures the antioxidant capacity of less polar (lipophilic) compounds compared to ABTS, which measures both hydrophilic and lipophilic compounds. This finding agrees with the reports of Sumaya-Martínez et al. [15].

As mentioned before, the differences between ABTS and DPPH values shown in Table 1 and those reported by other authors may be due to environmental conditions, harvest stage, agronomic practices, genetic base, and location, among others [2,4].

3.3. Optimization of the Extraction of PC and FC of Mango Peel Powder

3.3.1. Effect of Sequential Extractions

Figure 1 presents the cumulative percentages of PC and FC in the consecutive extractions.

In the case of PC, with only one extraction, 90.9% (3.78 mg GAE/g_extract) was achieved, in the second extraction, the cumulative percentage of compounds was 99.7%, and in the third extraction, 100% of compounds were extracted. The above coincides with the findings of Sánchez-Camargo et al. [30], which indicate that with two extractions, 100% of PC can be extracted. For FC, in the first extraction, 60.08% (1.37 mg QE/g_extract) was extracted, in the second extraction, an accumulation of 78.57% of FC was obtained, and in the third, 88.45%; upon performing the fourth and fifth extractions, 95.29% and 100% of FC were obtained, respectively. Based on this result, it was decided to use two sequential extractions for subsequent analysis, as they would represent 99.7% of PC and 78.57% of the FC present in the peel of mango var. “Kent”. This decision was taken because even though an additional extraction would increase the FC to 88.45%, it would have no effect on FC, giving, as a result, a more diluted extract.

3.3.2. Box–Behnken Design

Once the number of consecutive extractions needed to obtain the highest concentration of PC and FC was identified, a Box–Behnken design was used to determine the temperature, aqueous ethanol concentration, and extraction time for the maximum extraction of PC and FC.

Figure 2 presents the main effects plots for PC (mg GAE/g_extract) and FC (mg QE/g_extract). PC yield is affected by three factors (temperature, ethanol concentration, and extraction time); higher temperature and extraction time result in greater PC extraction (Figure 2a). Conversely, FC yield is influenced mainly by solvent concentration (ethanol), with higher solvent concentration leading to the greater extraction of flavonoids (Figure 2b).

It has been observed that the three factors (temperature, solvent, and extraction time) used in the design affect the extraction of PC and FC in different ways. Therefore, when performing an optimization to maximize the concentration of these compounds, it was decided that the extraction should be performed at 30 °C with ethanol (80%) for 90 min.

The desirability between both compounds (phenolics and flavonoids) was 0.8247 (Figure 3). Therefore, these conditions were selected as optimal for extracting the highest amounts of PC and FC.

3.4. Microencapsulation through SD and SFB

The microencapsulation achieved through SD had a process yield of 75.21%, whereas the yield through SFB was 34.92%. This is because for short operating times, with SFB, a significant portion of the solids remain adhered to the inert particles.

3.4.1. Retention and Encapsulation Efficiency

The retention and encapsulation percentages of PC of microcapsules produced by both drying methods are presented in Figure 4.

SD showed a 96.68% retention of PC and 92.44% encapsulation of PC. SFB showed a 98.28% retention of PC and 97.04% encapsulation of PC. The results show that the majority of the retained PC were encapsulated. There is no significant difference (p ≤ 0.05) in the retention and encapsulation efficiency with both drying methods.

3.4.2. Contents of BC and AC in Microcapsules

The contents of BC and AC in microcapsules produced through SD and SFB are presented in

Table 2. Contents of BC and AC in microcapsules obtained through SD and SFB.

Parameter	SD	SFB
PC (mg GAE/gdw)	102.35 ± 6.83 a	105.54 ± 4.34 a
FC (mg QE/gdw)	58.77 ± 0.42 b	67.56 ± 1.84 a
MC (mg ME/gdw)	7.26 ± 1.29 b	10.97 ± 0.45 a
AC (mg cya-3-glu/gdw)	0.064 ± 0.007 a	0.064 ± 0.005 a
CC (mg β-carotene/gdw)	0.11 ± 0.004 b	0.16 ± 0.002 a
AAC (mg AA/gdw)	0.95 ± 0.17 b	1.47 ± 0.02 a
Antioxidant capacity
ABTS (μmol TE/gdw)	141.34 ± 9.96 b,B	170.54 ± 4.36 a,B
DPPH (μmol TE/gdw)	268.15 ± 2.98 b,A	297.92 ± 6.03 a,A

Results are expressed as the mean ± standard deviation. Different lower-case letters per row indicate a significant difference (p ≤ 0.05). Different capital letters per column indicate a significant difference (p ≤ 0.05) in antioxidant capacity.

. The microcapsules obtained through SD and SFB presented no significant difference (p ≤ 0.05) regarding PC (mg GAE/g_dw) and AC (mg cya-3-glu/g_dw). The AC in both drying methods was 0.064 mg cya-3-glu/g_dw, which was very low and could be attributed to the high inlet air temperatures used in both drying processes (SD: 170 °C and SFB: 160 °C) since, as reported by Pajaro-Castro [31], high temperatures result in the degradation of anthocyanins.

Microcapsules obtained by SD and SFB presented a significant difference (p ≤ 0.05) of FC, MC, CC, AA, ABTS, and DPPH. The values of these phytochemicals were higher in the microcapsules obtained by SFB, as can be seen in Table 2. This could be attributed to the lower temperature conditions in the SFB process (160 °C inlet temperature) compared to the SD process (170 °C inlet temperature) and the more compact and thicker wall microcapsules obtained via SFB.

Regarding antioxidant capacity, Table 2 shows that the microcapsules obtained through SFB had significantly higher values of antioxidant capacity for ABTS and DPPH compared to microcapsules obtained through SD. These results agree with the higher content of BC present in SFB microcapsules (Table 1).

The microcapsules obtained through SD and SFB presented higher antioxidant capacity with DPPH than those obtained by ABTS. This could be attributed to the fact that mango peel contains a higher level of lipophilic antioxidants, as the DPPH method determines the antioxidant capacity of less polar (lipophilic) compounds, which agrees with the reports of Sumaya-Martínez et al. [15].

3.4.3. Moisture Content and a_w

Moisture content and a_w are presented in

Table 3. Moisture percentage and aw of the SD and SFB microcapsules.

Microcapsules	Moisture Content (%)	Water Activity (aw)
Spray Drying	3.70 ± 0.33 a	0.24 ± 0.01 a
Spout-Fluid Bed Drying	2.70 ± 0.26 b	0.16 ± 0.01 b

Data shown represent means ± standard deviation. Different letters per column show a significant difference (p ≤ 0.05).

.

The microcapsules obtained by SD had a moisture content significantly higher than that for SFB microcapsules (3.70 ± 0.33% compared to 2.70 ± 0.26%). These microcapsule moisture content results agree with those in other studies [4,32].

Regarding water activity, microcapsules obtained through SFB presented significantly lower values (0.16 ± 0.01) than SD microcapsules (0.24 ± 0.01). Water activity should be lower than 0.6, which suggests that the microcapsules are stable at both chemical and microbiological levels [32]. Based on the above, microcapsules obtained through both SD and SFB will exhibit good stability.

3.4.4. Particle Size Distribution

The distribution for SD is unimodal, while SFB shows a bimodal distribution (Figure 5). A difference in particle size and particle size distribution between SD and SFB was expected, due to their different drying mechanisms. In SD, the sprayed droplets are dried with hot air, primarily producing shrunken spheres. In SFB, the drying solution is sprayed onto a bed of inert solids; the fed solution forms a layer on the surface of the inert particles, and as it dries and becomes brittle, it is released from the surface of the inert particles, due to collisions and friction between the inert particles and the dryer wall. Therefore, the microcapsules obtained through SFB have irregular flake-like shapes with a larger and more varied particle size compared to those obtained by SD. According to Nikolić et al. [33], microcapsules with a larger particle size exhibit better performance in the delayed release of bioactive compounds.

Both drying methods resulted in a particle size ranging from 1.2 to 80 μm. The Sauter diameter (D[3,2]) was 10.35 ± 0.14 μm for SD and 10.84 ± 0.1 μm for SFB. The equivalent sphere diameter (D[4,3]) was 15.01 ± 0.06 μm for SD and 18.17 ± 0.36 μm for SFB.

3.4.5. Morphology of Microcapsules

Images of the microparticles obtained by SD and SFB using scanning electron microscopy (SEM) are presented in Figure 6. Most of the microcapsules obtained by SD exhibited a spherical shape with a smooth surface, varying in size; some of them were shrunken spheres with concavities (Figure 6a,b). This shape is typical of microencapsulated particles obtained through SD, due to the rapid evaporation of water [24]. Conversely, the microcapsules obtained by SFB have a more compact and thicker flake-like or sheet-like shape (Figure 6c,d), due to the mechanism by which this type of dryer operates (the release of broken dried layers from the surface of inert particles). The differences in shape and size of the microcapsules will result in different rates of release of BC during in vitro gastrointestinal digestion.

3.5. In Vitro Gastrointestinal Digestion

In vitro digestion process was developed on both unencapsulated extract and microencapsulated extract using SD and SFB methods. Figure 7 presents the percentage of recovery, %R (oral and gastric phases), and the percentage of bioaccessibility, %B (intestinal phase), of BC and AC of the unencapsulated extract, microcapsules through SD, and microcapsules by SFB. The percentages of BC and AC refer to the initial amounts in each sample before in vitro digestion.

Regarding phenolic compounds, Figure 7a shows that after the oral phase, only 86.4% of the phenolic compounds were found in the oral phase for the unencapsulated extract, whereas no phenolic compounds were detected for both microcapsules, which means that they are protecting these compounds in this phase. The %R and %B of the phenolic compounds for unencapsulated extract, microcapsules by SD, and microcapsules by SFB in the gastric phase show that there is a significant difference (p ≤ 0.05) in %R between the unencapsulated extract (73.28%), microcapsules by SD (25.21%), and microcapsules by SFB (25.54%), indicating that both forms of microcapsules release only a fraction of the PC that has been encapsulated; therefore, they are protecting most of these compounds from the gastric digestion process.

In the intestinal phase, Figure 7a shows that the PC released from the microcapsules by both SD and SFB (%B) increased considerably compared to the microcapsules found in the gastric phase, indicating that the phenolic compounds were protected during digestion and were available for possible absorption. Microcapsules produced by SFB showed significantly (p ≤ 0.05) higher bioaccessibility (86.63%) of PC compared to microcapsules produced by SD (61.95%). This may be attributed to the fact that microcapsules produced by SFB have larger particle sizes, a thicker wall, and are more compact than those produced by SD. Microcapsules with higher particle sizes exhibit better performance in the delayed release of phenolic compounds [33].

The considerable reduction in phenolic compounds from the gastric (73.28%) to the intestinal phase (19.99%) for the unencapsulated extract may be attributed to the instability of phenolic compounds, which is in accordance with the work of Ketnawa et al. [34], who indicated that the enzymatic hydrolysis of PC and aromatic molecules leads to the formation of derivatives (quinones or chalcones), which show instability at a high pH.

Blancas-Benitez et al. [35] reported a bioaccessibility of PC of 40.53% and 59.54% in the peel of mango var. “Ataulfo”, and, similar to the present study, they also reported a decrease in phenolic compounds.

Figure 7b shows the %R and %B of flavonoids in the unencapsulated extract, in microcapsules by SD, and in microcapsules by SFB, where the same trend as that observed for phenolic compounds was noticed. In the gastric phase, the %R of the unencapsulated extract was notably high (70.7%) whereas the microcapsules showed lower %R (SD = 45.08%; SFB = 22.65%), since not all flavonoids were released, protecting part of them from the gastric digestion process.

In the intestinal phase, conversely, the flavonoids in the unencapsulated extract decreased drastically to 10.44%, but those of the microcapsules increased to 53.84% for microcapsules produced by SD and to 68.85% for microcapsules produced by SFB, showing again that the microcapsules were able to protect the flavonoids, releasing them in the intestine for possible absorption. The decrease in flavonoids in the unencapsulated extract in the intestinal phase may be attributed to the fact that flavonoids, when not protected by a wall material, undergo degradation due to various conditions (enzymes, pH, temperature, or agitation) in the in vitro digestion process. Zheng et al. [36] reported that quercetin degrades after the in vitro digestion process, due to the conditions to which it is subjected (pH, temperature, enzymes, and agitation).

Microcapsules manufactured by SFB showed a significantly (p ≤ 0.05) higher bioaccessibility (68.85%) of flavonoids compared to microcapsules manufactured by SD (53.84%). This may be due, as explained before, to the larger particle size, thicker wall, and more compact SFB microcapsules compared to those manufactured by SD.

Figure 7c,g,h show the %R and %B of mangiferin and the antioxidant capacity measured with ABTS and DPPH, respectively, of the unencapsulated extract, microcapsules by SD, and microcapsules by SFB. As can be seen in these figures, a similar behavior to that described for phenolic compounds and flavonoids is observed. That is, in the gastric phase, the %R in the unencapsulated extract was significantly high (76.24% for mangiferin and 77.80% for antioxidant capacity, measured with ABTS, and 40.96% for antioxidant capacity, measured with DPPH) than the %R of SD microcapsules (45.04% for mangiferin, 23.28% for antioxidant capacity by ABTS, and 25.13% for antioxidant capacity, measured with DPPH) and SFB microcapsules (24.61% for mangiferin and 24.10% for antioxidant capacity, measured with ABTS, and 11.81% for antioxidant capacity, measured with DPPH), but after the intestinal phase, the %B of mangiferin and the antioxidant capacity, measured through both ABTS and DPPH, deceased drastically in the unencapsulated extract (8.21% for mangiferin and 8.71% for antioxidant capacity, measured with ABTS, and 7.88% for antioxidant capacity, measured with DPPH) compared to those for the SD microcapsules (99.95% for mangiferin and 47.31% for antioxidant capacity, measured with ABTS, and 52.73% for antioxidant capacity, measured with DPPH) and SFB microcapsules (69.98% for mangiferin and 72.60% for antioxidant capacity, measured with ABTS, and 75.74% for antioxidant capacity, measured with DPPH).

The microcapsules produced by SD showed a significantly (p ≤ 0.05) higher bioaccessibility (99.95%) of mangiferin compared to microcapsules produced by SFB (69.98%); however, regarding ABTS and DPPH, microcapsules produced by SFB showed significantly (p ≤ 0.05) higher bioaccessibility (72.60% and 75.74% for ABTS and DPPH, respectively) compared to microcapsules by SD (47.31% and 52.73% for ABTS and DPPH, respectively).

The results shown in Figure 7a–c,g,h (phenolic compounds, flavonoids, and mangiferin, measured with ABTS and DPPH), demonstrate that microencapsulation (SD, SFB) helped to avoid the degradation of BC and AC, yielding a high %B of these components after in vitro digestion. With the exception of mangiferin, the SFB microcapsules showed a better percentage of bioaccessibility than the SD microcapsules.

Conversely, regarding the %R and %B of anthocyanins, carotenoids, and ascorbic acid, Figure 7d,e show that, as seen with the unencapsulated extract, none of the microcapsules were able to protect these compounds during in vitro gastrointestinal digestion since, after the intestinal phase, the %B of these compounds were too small or undetectable. Figure 7d,e show that although a high %R for these compounds was obtained in the gastric phase (the microcapsules released a large amount of these compounds), after the intestinal phase, the %B was negligible or very small. Although anthocyanins are resistant to acidic media [37], they are unstable at a higher pH (≥6).

Regarding carotenoids, the %R depends on various factors that interfere with the digestive process, such as enzymes (α-amylase, pepsin, and pancreatin), pH (2–2.5), and temperature (37 °C), which can influence the release of carotenoids. Additionally, the release of carotenoids will depend on interactions with other compounds, like proteins, carbohydrates, lipids, or minerals [38]. The significant decrease (p ≤ 0.05) in the %B of carotenoids at the end of the intestinal phase could be attributed to the fact that carotenoids easily degrade when exposed to a high pH (pH ≥ 7), light, or heat. The %R and %B for carotenoids obtained in this study coincide with those reported by Cabezas-Terán et al. [39], who obtained %R levels of 68–102% and %B levels of 7.9–13.4% in mango byproduct microcapsules.

With respect to ascorbic acid, this compound is more stable in an acidic medium than in an alkaline one. In the gastric phase, HCl lowered the pH to 2.0, which helped stabilize the AA. The decrease in AA in the intestinal phase could be due to changes in pH (7), temperature, and enzyme action, which can contribute to the oxidation (degradation) of AA. The sensitivity of AA to conditions in the small intestine has previously been reported [40].

The results shown in Figure 7 demonstrate that for bioactive compounds that are stable at pH 7, the microencapsulation process (using both drying methods) adequately protected them from in vitro digestion. Likewise, microencapsulation (via SD or SFB) retained the antioxidant capacity (established with ABTS or DPPH) of the bioactive compounds, compared to the non-microencapsulated sample. The function of the microcapsules is to protect the CB and release them in the small intestine as a compound can only exert benefits if it is available for absorption after all the phases involved in the gastrointestinal digestion process have taken place [40].

4. Conclusions

“Kent” variety mango peel is a good source of bioactive compounds with an antioxidant capacity that can provide health benefits to humans; therefore, it is important to extract these valuable compounds. The optimal conditions for extracting bioactive compounds from “Kent” mango peel were: two extractions at 30 °C over 90 min with 80% ethanol. After in vitro digestion, the non-microencapsulated extract showed low or no bioaccessibility of CB and AC, while the microcapsules (manufactured via SD or SFB) were able to protect the BC (phenolic compounds, flavonoids, and mangiferin). Likewise, the microcapsules were able to retain the antioxidant capacity (measured with ABTS or DPPH) of the CB. In general, microcapsules produced by SFB better protected the bioactive compounds and, therefore, showed a better bioaccessibility of antioxidant capacity compared to microcapsules produced by SD.

Microcapsules obtained from the peel of mango var. “Kent” can be used in the food industry as an ingredient to produce functional foods and, thereby, to obtain the benefits of the compounds present in this fruit. More studies should be carried out to verify the biological activity of the microcapsules.