1. Introduction
The confectionery industry continually seeks innovative approaches to find colouring molecules that can compete with synthetic analogues, enhancing product appeal while meeting consumer demands for healthier options [
1]. Another strategy in the food industry is to replace food additives by removing E codes from labels and using natural food ingredients with colouring properties. These strategies utilize less processed vegetable materials to achieve clean label solutions, maintaining food colour, quality, and safety. Due to these changes, the food industry aims to replace synthetic dyes such as Brilliant Blue (E133) and Allura Red (E129) with natural dyes that possess bioactive properties [
2]. These natural dyes can be derived from leaf and peel co-products of the agri-food industry, transforming waste into valuable resources.
Anthocyanins are a group of water-soluble pigments responsible for the colours ranging from red to purple to blue in many fruits, vegetables, flowers, and leaves. The colours produced by anthocyanins have been studied for several factors, including light [
3], temperature [
4], pH [
3,
5], copigments [
3,
6,
7,
8], metal ions, and other environmental factors.
Some studies [
3,
6,
7] have suggested that anthocyanin degradation is accelerated as temperature increases, and the loss of colour is more pronounced in the presence of phenolic acids and other flavonoids, such as rutin [
3]. However, the stability of anthocyanins can be increased through copigmentation, which involves the association between anthocyanins and one or more flavonols through hydrogen bonds. This copigmentation forms a protective structure resistant to external factors such as pH. Other authors [
3] have noted that in acidic conditions, anthocyanins tend to appear red, while in more neutral conditions, they can appear purple or blue [
3]. The colour variation in anthocyanins with pH is due to the equilibrium of molecular species in solution as pointed out by [
9].
The structures of anthocyanins can vary depending on the specific compound and the plant species from which they come. However, they typically consist of anthocyanidins derived from flavonoids. Anthocyanins are often glycosylated, meaning they have sugar molecules attached to the aglycone. These sugar groups can include glucose, galactose, arabinose, rhamnose, or other sugar molecules [
3]. Some anthocyanins may also have acyl groups attached to the sugar molecules, such as acetyl, coumaroyl, caffeoyl, or organic acids (malonic, succinic, tartaric, and malic acid) [
10]. Acylated anthocyanins are more stable to pH, temperature, and interaction with other matrix compounds than non-acylated anthocyanins [
11].
Deoxyanthocyanidins (3-DXAs) are distinguished from anthocyanidins due to the absence of a hydroxyl group at position 3, an indication that at pH 1–7, no difference is observed in the colour of 3-DXAs extracted from sorghum [
5]. However, some authors claim that it is possible to extract the violet colour from sorghum when using an aqueous solution of pH 9 [
12]. In sorghum, the forms of 3-DXA are apigeninidin and luteolinidin, which give the yellow and red colour, respectively [
13,
14].
Several studies have been published on the extraction of these compounds using some solvents, such as alcohols [
15], alkaline water [
16], acidified water [
15,
16,
17], or ionic liquids [
18,
19,
20] to enhance the extraction efficiency. However, these methods are time-consuming, require larger volumes of solvents, are high-cost, and have potential environmental impacts. Other methods have been applied by other authors like ultrasound-assisted (UA) [
21], microwave-assisted (MA) [
22], and pulsed electric field (PEF) extraction [
23], they are distinguished by increasing the availability of anthocyanins in the extraction solvent, thus increasing the extraction yield. However, they require specialized equipment and careful control of electrical parameters [
16,
24,
25].
Apart from their role in plant pigmentation, anthocyanins have gained recognition in several studies for their antioxidant [
18], anti-inflammatory, and potential anti-cancer properties. Furthermore, acylated and non-acylated anthocyanins have been reported for prebiotic, antidiabetic, and antioxidant properties, with the consumption of plant sources with these molecules playing an important role in health [
11,
26,
27].
Edible coatings, commonly used in the food industry, are proteins, polysaccharides, or lipids. Generally, proteins are used to provide mechanical stability to the film, polysaccharides to control gas exchange, and lipids to reduce water transmission [
28]. The choice of polymeric materials influences the texture of the films, giving shine or opacity, extending shelf life, enhancing sensory attributes, and improving nutritional value [
29,
30]. Integrating anthocyanin-rich extracts into edible coatings offers several properties by the formation of a uniform film to enhance the adhesion and stability of anthocyanins during the industrial process [
28,
31,
32]. These coatings provide a protective barrier against moisture and light transmission, and act as an encapsulating agent, immobilizing anthocyanins and protecting them from light, heat, pH changes, and oxidation [
3,
29].
A crucial aspect of evaluating anthocyanin-based edible coatings is their temperature resistance. Arabic gum is one of the polymeric materials most used in coatings in the confectionery industry, due to its solubility in water and ability to form gels [
33]. Locust gum, used in the food industry as a thickening, stabilizing and gelling agent, or emulsifier (E410), is more resistant to temperature than Arabic gum [
34] and has antioxidant capacity [
35]. This study aims to evaluate the resilience of these two coatings to temperature variations and their role in stabilizing anthocyanin-based dyes applied to sugar syrup, in terms of colour discolouration due to sugar syrup.
Another critical consideration in utilizing anthocyanin-based coatings is their sensitivity to pH values [
31]. Obtaining a stable purple hue directly from anthocyanin- or 3-DXA-based plant matrices, without mixing blue and red dyes, would simplify the formulation process and offer more consistent colours throughout the product’s shelf-life.
Beyond their visual impact, the biocapacity of anthocyanins is a key attribute to their appeal in confectionery applications. Incorporating anthocyanin-rich coatings into confectionery products offers the potential to deliver antioxidant benefits to consumers. However, ensuring the retention of bioactive compounds throughout processing and storage poses a challenge.
This study intends to obtain a stable purple colour directly from anthocyanin-based plant matrices, specifically using leaves and peels from agri-food by-products, investigate the stabilization of anthocyanin-based dyes using Arabic and locust gum coatings, and evaluate the resilience of these coatings to temperature variations in terms of colour discolouration when applied to sugar syrup.
2. Materials and Methods
2.1. Chemicals and Materials
Ethanol p.a., citric acid, sodium citrate, gallic acid, hydrochloric acid, ascorbic acid, iron (II) sulphate heptahydrate, iron (III) chloride hexahydrate, and sodium acetate trihydrate were purchased from Merck (Darmstadt, Germany). Sodium carbonate anhydrous, sodium bicarbonate, Folin–Ciocalteu reagent, potassium hexacyanoferrate (III), and anhydrous sodium sulphate were purchased from Panreac (Barcelona, Spain). 2,4,6-tris (2-pyridyl)-S-triazine (TPTZ) and ferric chloride were acquired from Fluka (Buchs, Germany). Ethanol absolute anhydrous was purchased from Carlo Erba (Marseille, France). Other reagents such as potassium chloride, soluble potato, starch, sodium phosphate, 4-nitrophenyl-α-D-glucopyranoside, 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid (trolox), 2,2-diphenyl-1-picrylhydrazyl radical (DPPH•), 3,5-dinitrosalicylic acid (DNS), and the enzymes α-amylase from porcine pancreas and α-glucosidase from Saccharomyces cerevisiae were purchased from Sigma-Aldrich (Sternheim, Germany). Acarbose was purchased from Alfa Aesar (Karlsruhe, Germany). Food grade additives such as locust bean gum was purchased from Sosa (Ingredients S.L., Spain, and Arabic gum was purchased from Sigma-Aldrich (Darmstadt, Germany). The sugar (Sidul, Lisbon, Portugal) and the solid white sugar paste (Auchan) were purchased from a commercial store Auchan (Lisbon, Portugal). All other unlabelled chemicals and reagents were analytical or HPLC-MS Optima grade.
2.2. Plant Material
Fresh black carrots (Daucus carota ssp. sativus var. atrorubens), radishes (Raphanus sativus), and purple sweet potatoes (Ipomoea batatas (L.) Lam) were acquired from a commercial store, Celeiro, in Sintra, Portugal. Dried sorghum (Sorghum bicolour L.) was purchased from PMA28 in Varize, France, and ground in a mill (IKA Micro Fine Mill Culatti) using a 1.0 mm thick sieve. The powdered sorghum was stored under vacuum in a packaging film polymer (LDPE 60 µm/PA 30 µm) from Amcor Flexibles, Oeiras, Portugal, and placed in desiccators until further analysis.
2.3. Anthocyanins Extracts
Anthocyanins from black carrot, sweet potato (peel and pulp), and radish (peel) were extracted according to [
36] with modifications and 3-deoxyanthocyanidins from sorghum using the [
37] methodology. The material was peeled and cut into small pieces (2 cm
2), and mixtures with alkaline water at pH 8, adjusted with sodium hydrogen carbonate (0.103 mg mL
−1 NaHCO
3) and sodium carbonate (0.008 mg mL
−1 Na
2CO
3), were prepared with a solid–liquid ratio of 1:5 (
w/
v) for black carrots, and mixtures with acidified water at pH 3, adjusted with citric acid (0.961 mg mL-1, C
6H
8O
7), were prepared with a solid–liquid ratio of 1:3 (
w/
v) for radish, sweet potato peels, and 1:1 (
w/
v) for sweet potato pulp.
Anthocyanins from black carrots, radish, and purple sweet potato were extracted in two different conditions: 1) using an alkaline aqueous solution at pH 8, adjusted with sodium hydrogen carbonate (0.103 mg mL−1 NaHCO3) and sodium carbonate (0.008 mg mL−1 Na2CO3) (extracts at pH 8), and 2) using an acidified water at pH 3, adjusted with citric acid (0.961 mg mL−1, C6H8O7) (extracts at pH 3). In both cases, the extraction mixtures were heated on a hotplate (Heidolph MR 3001, Schwabach, Germany) for 10 min and homogenized using an ultra-turrax T25 (Janke & Kunkel, Staufen, Germany) at 8000 rpm for 10 min. Subsequently, all the extraction mixtures were sonicated (ultrasonic processor, Hielscher UP200S, Berlin, Germany) at 50% amplitude and 0.5 cycles for 10 min.
Sorghum 3-DXA were extracted from sorghum leaves using two extraction methods: solvent extraction and ultrasound-assisted (UA) at room temperature. In both cases, the ground plant material was mixed with an alkaline aqueous solution at pH 10 in a 1:20 (w/v) solid to liquid ratio. For the solvent extraction, the mixture was placed on an orbital shaker (Unitronic-OR, Selecta, Barcelona) at 25 °C and stirred at 70 rpm for 30 min. In the ultrasound method, the extraction mixtures were sonicated (ultrasonic processor, Hielscher UP200S, Berlin, Germany) at 50% amplitude and 0.5 cycles for 10 min.
After the extraction step, all the extracts were filtered through a 150 mm diameter filter (Whatman No. 1 Qualitative) at reduced pressure (GAST DOA-P104-BN, Redditch, United Kingdom). The supernatants were frozen at −80 °C, freeze-dried (Scanvac Cool Safe, Labogene, Bjarkesvej, Denmark). The vacuum pressure of the freeze-drier was set at 0.2 hPa, the plate temperature was 20 °C, and the condenser was at −50 °C for 24 h. The powdered colouring matters were weighed and stored in glass bottles (Schott 250 mL, Stuttgart, Germany) inside desiccators containing silica gel, kept in the dark at room temperature. All the extractions were performed in duplicate.
2.5. Characterization of the Plant Extracts
The extracts were characterized both chemically (structural characterization of anthocyanins and content of total anthocyanins and total phenolic compounds) and functionally (antioxidant and antidiabetic capacities). A commercial food colouring (dye factory) of unknown composition, used as a control in the formulation of colouring coatings, was also analysed.
2.5.1. Chemical Characterization of the Plant Extracts
Analysis by HPLC-DAD-MS and UHPLC-HRMS/MS
The identification and structural characterization of anthocyanins present in the extracts was achieved by high-performance liquid chromatography (HPLC) coupled with mass spectrometry (MS) assays.
Aliquots of 10 µL of each extract were initially analysed on a LC-MS system, consisting of an HPLC Ultimate 3000RS with Diode Array Detector (DAD) coupled to an ion trap LCQ Fleet mass spectrometer (Thermo Scientific, Carlsbad, CA, USA), with an ESI source. The full scan acquisition was performed in the positive ion mode in the range 100–1500 m/z. The DAD system was monitored in the range between 220 and 700 nm, and additional UV/VIS spectra were recorded at 520, 320, and 280 nm.
The identification and structural characterization of anthocyanins were achieved with an UHPLC Elute system coupled to a QqTOF Impact II high-resolution mass spectrometer (Bruker Daltonics, Bremen, Germany), interfaced with an ESI source operating in the positive mode. Mass spectra were acquired in the data-dependent mode (DDA) in a mass range between 100 and 1500 m/z, with an acquisition rate of 3 Hz, using a dynamic method with a fixed cycle time of 3 s.
The anthocyanins were separated by a Kinetex C18 core–shell column (150 mm × 2.1 mm; particle size 2.6 μm; Phenomenex, Torrance, CA, USA), using an elution gradient of 0.1%
v/
v formic acid in water (mobile phase A) and methanol (mobile phase B), at a flow rate of 300 μL min
−1, More experimental details can be found in [
13].
Total Antocyanin Content
The total anthocyanin content (TAC) was determined by the pH differential method following the methodology described by [
38]. This methodology involves measuring the absorbance of anthocyanin extracts at two different pH values (pH 1 and 4.5) and using these absorbance values to calculate the concentration of anthocyanins. Anthocyanin extracts were prepared at concentrations of 1.28 mg/mL for radish and sweet potato peel, 2.46 mg/mL for sweet potato pulp, and 0.63 mg/mL for black carrot, sorghum, and commercial colourant. The TAC was calculated using the equation:
where
A is the difference in absorbance between pH 1.0 and pH 4.5,
DF is the dilution factor,
MW is the molecular weight of cyanidin-3-glucoside (the anthocyanin standard 484.83 g mol
−1) or the specific anthocyanin being analysed (in sorghum apigenidin, 255.24 g mol
−1),
ε is the molar absorptivity coefficient of cyanidin-3-glucoside and apigenidin depending on the plant in study, and
l is the path length of the cuvette (1 cm).
Total Phenolic Compounds Content
The total phenolic content (TPC) of the extracts was estimated using the Folin–Ciocalteu colourimetric method, as described by [
39] and modified by [
40], with gallic acid as the standard phenolic compound. Diluted samples (1–3 mL) were added to 10 mL volumetric flasks containing distilled water and 0.5 mL of Folin–Ciocalteu reagent and shaken. After 5 min, 1.5 mL of a 200 g L
−1 sodium carbonate solution was added, and the volume was adjusted to 10 mL with distilled water, mixed, and left to stand for 2 h. A blank reagent with distilled water was prepared. Absorbance was measured at 750 nm using a double-beam UV–visible spectrophotometer (Hitachi U-2010, USA). Concentrations of total phenolic compounds were determined based on the standard curve (y = 88.003x + 0.0288, r
2 = 0.998) in terms of grams per liter of gallic acid equivalents (GAEs). The linearity range for this assay was 6.3 × 10
−4−1.3 × 10
−2 gL
−1 GAE, with an absorbance range of 0.08–1.13 AU. All determinations were performed in triplicate.
2.5.2. Biological Capacities of the Extracts
Antioxidant Capacity
The scavenging effect of the DPPH
• free radical was assessed spectrophotometrically by the modified method of [
41]. A volume of 0.1 mL of each plant extract at various concentrations was added to 2 mL of 0.07 mmol L
−1 DPPH
• in 950 g L
−1 ethanol, followed by shaking and incubation for 60 min at room temperature in the dark. The absorbance was measured at 517 nm using a double-beam UV–visible spectrophotometer (Hitachi U-2010, USA). A blank sample with ethanol and DPPH
• solution served as the negative control. Antioxidant capacity results were expressed as trolox equivalents (TEs) based on a standard curve (y = −0.0011x + 0.0109, r
2 = 0.999) with a linearity range of 23.97–800 µmol TE g
−1 and an absorbance range of 0.037–0.85 AU. All determinations were performed in triplicate.
Ferric ion reducing antioxidant power (FRAP) measures the formation of a blue-coloured Fe
2+-tripyridyltriazine compound from the rust-coloured oxidized Fe
3+ form by the action of electron-donating antioxidants. The FRAP assay was performed using the modified methodology of [
42]. FRAP reagent was prepared with 1 mmol L
−1 TPTZ and 2 mmol L
−1 ferric chloride in 0.25 mol L
−1 sodium acetate at pH 3.6. Diluted extracts (200 µL in 500 g L
−1 methanol) were mixed with 1.8 mL FRAP reagent, allowed to stand for 4 min at room temperature, and the absorbance of the blue complex was measured at 593 nm against a water blank using a double-beam UV–visible spectrophotometer (Hitachi U-2010, USA). A standard curve (y = 0.0213x + 0.0057, r
2 = 0.998) was prepared using different concentrations of iron sulphate, with a linearity range of 1.25–50 µmol L
−1 and an absorbance range of 0.02–1.03 AU. FRAP values are presented as µmol Fe
2+ g
−1 of sample. All determinations were performed in triplicate.
Antidiabetic Capacity
Antidiabetic capacity was assessed by evaluating the potential of the extracts to inhibit the carbohydrate digestive enzymes α-amylase and α-glucosidase, according to the procedures described by [
43,
44,
45], respectively.
For the α-amylase inhibition assay, radish, black carrot, and purple sweet potato extracts were dissolved in water (20 μg mL−1) and then diluted to concentrations ranging from 2.5 to 18 μg mL−1. Sorghum extract was dissolved in 50% (v/v) ethanol:water (10 μg mL−1) and diluted to concentrations ranging from 2 to 9 μg mL−1. For the assay, 100 µL of α-amylase (0.5 mg mL−1 in sodium phosphate buffer 0.02 M, pH 6.9, 6.7 mM NaCl) and 100 µL of each extract concentration were incubated at 37 °C for 10 min. Then, 100 µL of starch suspension (1% w/v in sodium phosphate buffer 0.02 M, pH 6.9, 6.7 mM NaCl) were added, and the mixtures were further incubated at 37 °C for 10 min. After that, 200 μL of 3,5-dinitrosalicylic acid reagent (20 mL of 96 mM DNS, 8 mL of 5.315 M sodium potassium tartrate tetrahydrate in 2 M NaOH, and 12 mL of water) were added, and the tubes were boiled on a heating block (Bioblock Scientific Code 92607) at 100 °C for 15 min. Afterwards, the mixtures were cooled in an ice bath, 2 mL of distilled water was added, and the absorbance was read at 530 nm on a UV-Vis spectrophotometer (SPEKOL 1500, Analytik, Jena, Germany). The mixtures without plant extracts were used as negative controls and mixtures without α-amylase were used as samples’ blanks. Increasing concentrations of acarbose (5.0–100.0 µgmL−1) were used as positive controls.
The enzyme inhibition rate was calculated according to Equation (3):
where
AbsCN is the absorbance of the negative control,
AbsA the absorbance of the samples, and
AbsB the absorbance of the sample blanks. Values were assessed in triplicate, and the results were expressed as the final concentration (mg mL
−1), in the reaction mixture, which reduces the enzyme activity by 50% (IC
50). Data are presented as means ± standard deviations.
For the α-glucosidase assay, the extracts were first dissolved in water to a concentration of 5 mg mL
−1 and then diluted to concentrations ranging from 0.2 to 2.0 mg mL
−1. The α-glucosidase (6.25 units mL
−1) and 4-nitrophenyl-α-D-glucopyranoside (2.5 mM) were prepared in 0.1 M sodium phosphate buffer at pH 6.9. Reaction mixtures containing 5 µL of α-glucosidase, 125 µL of phosphate buffer (pH 6.9, 0.1 M), and 20 µL of the different extract concentrations were prepared in a 96-well microplate (Greiner Bio-One, Rainbach im Mühlkreis, Austria) and incubated for 15 min at 37 °C. Then, 20 µL of 4-nitrophenyl-α-D-glucopyranoside was added, and the plates were incubated for an additional 15 min, at 37 °C. To stop the reactions, 80 µL of 0.2 M Na
2CO
3 was added in each well. The absorbance was measured in a microplate reader (FLUOstar
® Omega Plate Reader, BMG Labtech, Ortenberg, Germany) at 405 nm. Reaction mixtures without extracts were used as negative controls, and reaction mixtures without the enzyme were used as samples’ blanks. Increasing concentrations of acarbose (80–980 µg mL
−1) were used as positive controls. The extracts’ inhibitory capacity was calculated according to Equation (5):
where
AbsCN is the absorbance of the negative control,
AbsA is the absorbance of the sample, and
AbsB is the absorbance of the sample blanks. Values were assessed in quadruplicate, and the results were expressed as the final concentration (mg/mL) in the reaction mixture, which reduces the enzyme activity by 50% (IC
50). Data are presented as means ± standard deviations.
2.6. Confectionery Colouring Coatings
The methodology for the formulation of colouring coatings, based on anthocyanins and 3DXAs, to be applied to confectionery products, was adapted from the factory manufacturing process and involved the selection of purple colouring extracts (radish, black carrot, purple sweet potato, and sorghum) and polymeric materials (locust gum and Arabic gum) for application in sugar syrup. The concentrations of the colouring materials were adjusted according to the compatibility and desired characteristics of the chocolate almonds (
Table 1). The control was a commercial colourant (Dye factory) of unknown composition. The quantity of colouring materials was selected, based on the same absorption (Abs = 1), at 510 nm for anthocyanins and 420 nm for 3-DXAs, in scanning the UV-Vis spectrum. The colouring materials were added to the polymeric materials and to the sugar syrup at 50 °C. The solutions were homogenized in the ultra-turrax (T25, Janke & Kunkel) at 8000 rpm for 5 min.
2.7. Thermostability of Confectionery Colouring Coatings
The stability of the formulated colouring materials was evaluated at a temperature of 90 ± 5 °C for 75 min. Samples (7 mL) of the formulations indicated in
Table 1 were placed into test tubes (20 mL) and sealed hermetically. A commercial dye was used as a control in the temperature stability tests. The tubes were placed in the oven (Ventilcell 111, MMM Group), and duplicates were removed every 15 min and immediately cooled in an ice bath to stop thermal degradation. The changes in the total anthocyanins and 3-DXA contents during heat treatment were determined by measuring the absorbance at 510 and 520 nm, respectively, using a UV–visible spectrophotometer (dual beam; Hitachi U-2010). Assays were performed in duplicates.
The absorbance measurements of the colouring coating formulations were used to obtain the kinetic parameters according to [
13] to estimate the thermostability of the anthocyanins and the 3-DXAs. The first-order reaction rate constants (K
d), half-lives (t
1/2), i.e., the time that is necessary for degradation of 50% of anthocyanins and 3-DXAs, were calculated by the following Equations (5) and (6):
where
C0 is the initial concentration of the anthocyanins or 3-DXAs, and
Ct was the anthocyanins or 3-DXAs concentration after t minutes at 90 °C.
The integrated rate equation for a second-order reaction is determined from experimental data. It can be calculated from the initial rate of reaction and the initial concentrations of the anthocyanins or 3-DXAs. The rate equation is represented as:
where
k is the rate constant and [
A]
t is the concentration of the anthocyanins or 3-DXAs after t minutes of heating at 90 °C, and [
A]
0 is the initial concentration of the anthocyanins or 3-DXAs.
2.8. Purple Confectionery Coatings: Effects of Blue Dye Addition
The methodology for the formulation of purple-coloured polymeric coatings, was adapted from the factory manufacturing process, and incorporating anthocyanins and 3-DXAs extracts, were developed with and without the addition of spirulina as a blue dye (
Table 1 and
Table 2, respectively). These coatings were meticulously applied onto white solid sugar pastes, each cut into 5 cm
2 squares, following the procedure outlined in
Section 2.6. The application involved brushing the pastes three times at 30 min intervals, followed by drying in an oven set at 30 ± 5 °C. Subsequently, the samples underwent a 72 h drying period under the same temperature conditions.
2.9. Colourimetry
The colour of the coating pastes was assessed using colourimetry and compared against purple Easter almonds dyed with clean label colourants and those with synthetic dyes. All experiments were conducted in duplicate.
The colourimetric analysis was performed according to [
13]. The colour properties in the CIELAB system, including lightness (L*), red-green chromaticity (a*), and yellow-blue chromaticity (b*), were measured in liquid samples using a Chroma Meter CR-400 (Konica Minolta, Japan), with water as the standard reference. For solid samples, colour measurements were determined with a CS-5 CHROMA SENSOR spectrophotometer (Datacolour International) using a 45/0 geometry, D65 illuminant, 10° angle, and including the specular component, with the CIELAB colour white as the standard reference. The colour difference between the samples was calculated using the equation:
where ΔL*, Δa*, Δb* represent the difference between each parameter for the anthocyanin and 3-DXA colourants. The measurements were performed in triplicate.
2.10. Statistical Analysis
The results were submitted to one-way analysis of variance (ANOVA) using multiple comparison tests (Tukey HSD) to identify differences between groups. Statistical analyses were tested at a 0.05 level of probability. The range, mean, and relative standard deviation (RSD) of each parameter were calculated using the software, StatisticaTM 12.0 [
46].
4. Conclusions
The extraction at acidic pH yields higher total anthocyanin content for most plant extracts, except for black carrot and sorghum, while extraction at alkaline pH produces a more purple colour but less stable anthocyanins.
The total anthocyanin content and total phenolic content of plant extracts varied significantly, with ultrasound-assisted extraction at acidic pH yielding the highest values for radish extracts.
The plant extracts exhibited significant antioxidant capacities and enzyme inhibitory capacities, with variations influenced by the type of assay and polyphenol composition, indicating that sorghum extract, in particular, holds potential for managing postprandial hyperglycemia and type 2 diabetes.
The UV-VIS spectra obtained from HPLC-DAD conjugated with tandem mass spectrometry data, in positive ESI mode, demonstrated that the majority of anthocyanins in the extracts are acetylated, with the exception of sorghum, which contains apigeninidin, a primary pigment belonging to the 3-DXA group.
HPLC clearly demonstrated quantitative, but not qualitative, differences between the extracts of the peel and pulp of sweet potatoes. Thirteen anthocyanins, consisting of non-acylated or acylated peonidin and cyanidin glycosides, were identified. The major anthocyanin is a peonidin 3-caffeoyl-p-hydroxybenzoyl sophoroside-5-glucoside.
Extracts of black carrot contained five cyanidin-3-xyloslyl-glucosyl-galactosides, three of which are mono-acylated with ferulic, sinapic, and p-coumaric acids. The variety analysed also had traces of acylated peonidin and pelargonidin anthocyanins.
The extract of radish peel predominantly contained pelargonidin derivatives, as indicated by a blue shift in UV-VIS spectra. Sixteen pelargonidin-based compounds were identified, with the presence of one or two cinnamoyl groups on the di-glycosylated fraction at the C3 position and a malonyl at hexose at the C5 position.
The extract of dye factory showed cyanidin and peonidin acylated derivatives, with fragmentation patterns suggesting the presence of anthocyanin-3-(cinnamoyl) sophoroside-5-glucoside derivatives, similar to those in the peel and pulp of purple sweet potato extracts.
Locust bean gum provided better temperature resistance and lower degradation percentages for coloured coatings of plant extracts compared to Arabic gum, making it more effective for stabilizing anthocyanins during thermal processing.
Thermostability tests revealed that sorghum extracts exhibited the highest colour variation when coated with gums, while black carrot extracts coated with Arabic gum and dye factory extracts coated with locust bean gum showed the best colour stability, with locust bean gum providing comparable stabilization to extracts without any coating.
In sugar pastes, sweet potato pulp extracts exhibited less colour variation than dye factory and synthetic dyes when used alone and, when combined with spirulina blue dye, provided a more stable and subtle colour variation in polymeric-coated sugar pastes, making it a superior and more consistent choice for colouring almonds purple.
Future work could investigate whether anthocyanin-based coatings retain their biocapacity when applied to confectionery products. Conclusions may include findings on the ability of coatings to preserve antioxidant levels and potential health benefits in final products.