**Stability Analysis of Anthocyanins Using Alcoholic Extracts from Black Carrot (***Daucus Carota* **ssp.** *Sativus* **Var.** *Atrorubens* **Alef.)**

**Guillermo Espinosa-Acosta 1 , Ana L. Ramos-Jacques 2 , Gustavo A. Molina 1 , Jose Maya-Cornejo 3 , Rodrigo Esparza 3 , Angel R. Hernandez-Martinez 3 , Itari Sánchez-González <sup>4</sup> and Miriam Estevez 3, \***


Received: 29 August 2018; Accepted: 22 October 2018; Published: 24 October 2018

**Abstract:** Anthocyanins are used for food coloring due their low toxicity and health benefits. They are extracted from different sources, but black carrot has higher anthocyanin content compared with common fruits and vegetables. Here, we study alcoholic anthocyanin extracts from black carrot to enhance their stability. The objective of our research is to determine if microencapsulation with tetraethyl orthosilicate (TEOS) is a feasible option for preventing black carrot anthocyanin degradation. Extraction solvents were solutions of (1) ethanol/acetic acid and (2) ethanol/citric acid. Samples were purified through a resin column and microencapsulated using TEOS. Fourier Transformed Infrared Spectroscopy (FTIR) spectra of samples were obtained, and degradation studies were performed under different conditions of UV radiation, pH and temperature. Antioxidant activity was evaluated with radical 2,2-diphenyl-1-picrylhydrazyl (DPPH) scavenging and electrochemical cupric reducing antioxidant capacity (CUPRAC). Color evaluation on food models were performed with CIE Lab at the beginning of experiments and after 25 days of storage. Results indicate that the more stable extracts against pH media changes are samples obtained with ethanol/acetic acid solution as extraction solvent. Extract purification through resin and TEOS microencapsulation had no significant effect on extract stability. In conclusion, although TEOS microencapsulation has proven to be effective for some dried materials from natural extracts in our previous research, we do not recommend its use for black carrot extracts considering our results in this particular case.

**Keywords:** anthocyanin; natural extract; tetraethyl orthosilicate; black carrot; antioxidant activity

#### **1. Introduction**

The use of extracts from natural sources as food coloring is an ongoing trend because, in general, they are Generally Recognized as Safe (GRAS) substances and bring health benefits for consumers [1]. A variety of natural colorants are used in the food industry, but there is still a concern about their production costs and stability and their performance had been studied for the past decade [2–6]. Substances like carotenoid, chlorophyll, turmeric, anthocyanin, and betalain extracts from natural sources impart a variety of colors [7,8].

Our hypothesis relies on the possibility of using microencapsulation (with TEOS) to increase stability of anthocyanins. Previously, we studied betalain extracts from *Beta vulgaris* and *Myrtillocactus geometrizans* and obtained dried materials that were microencapsulated using TEOS, obtaining an improvement in the materials' stability against UV light, pH and temperature [1].

Anthocyanins (polyhydroxy and polymethoxy derivatives of 2-phenylbenzopyrylium of flavylium salts) are a group of phenolic compounds that are responsible for the colors of flowers, fruits, and vegetables [9]. It has been reported that this group of compounds has antioxidant [10], antimutagenic, anticancer and antiobesity properties, and they reduce the risk of coronary heart disease [9,11,12]. The colors imparted by anthocyanins are bright and could be used in the food industry as a replacement for colorants like FD&C Red 40 [13]. The colors obtained are commonly red, orange, blue and purple, depending on the chemical structure of anthocyanins; but, structural transformations are induced by changes in the pH of the medium, that affect both color quality and intensity [14].

Common sources of anthocyanins are purple corn, red cabbage, purple sweet potato, apples, grapes, kiwi, red onions and several berries [6,15]. Some studies have used local plants like black carrot [6]. Black or purple carrot (*Daucus carota* ssp. *sativus* var. *atrorubens* Alef.) was originally from Turkey and the Middle and Far East [16], but recently, new varieties with high anthocyanin content had been cultivated in other parts of the world [12,13,16]. For those reasons, this source was selected for our study.

The stability of anthocyanins in plant extracts depends on the temperature and solids content; and by increasing these conditions, the degradation rates of anthocyanins increase too. To minimize this degradation, it is recommended to cool the concentrates as soon as produced. Other factors that affect color and stability of anthocyanins are concentration, light, presence of co-pigments, metallic ions, enzymes, sugars, proteins, and antiradical activity (which quantifies the ability of complex chemical structures to scavenge free radicals) [2,4,17–21].

Considering those factors, many efforts have been made to enhance anthocyanins' stability. Acylation of anthocyanins is the most commonly used method, as it has been reported that acylated cyanidin derivatives are more stable during prolonged storage compared to the corresponding non-acylated ones [12,13,16] Stintzing et al. [10] also confirmed that there is an increase in color strength through acylation. On the other hand, microencapsulation has been used to increase stability of natural colorants (anthocyanin and betalain derivatives) [2,4,17–19]. Then, considering these conditions, some efforts have been made to preserve natural extracts and colorants using different techniques, such as removing compounds through resin columns and microencapsulation with TEOS.

Here, we study the stability of black carrot extracts while modifying the experimental conditions, including extraction solvents, and we study the feasibility of microencapsulation with TEOS and the repercussion of this procedure on the stability of the extracts.

#### **2. Results and Discussion**

### *2.1. Anthocyanin Content*

Major groups of substances were quantified with an UV-Vis spectroscopy analytic method. Figure 1 shows the Fourier Transformed Infrared (FTIR) spectra of samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA) and the IR spectra obtained by simulation. In Figure 1a, vibrations obtained for both samples showed a band at 980 cm−<sup>1</sup> of an C-H in plane deformation, at 1070 cm−<sup>1</sup> , corresponding to an aromatic ring C-H deformation, bands at 1620 and 1447 cm−<sup>1</sup> , that correspond to vibration (C=O) of the benzopyran aromatic ring and 1590 cm−<sup>1</sup> from the stretching vibration (C=C) of an aromatic ring, a band at 1235 cm−<sup>1</sup> , that corresponds to stretching of pyran rings, typical of flavonoid compounds, and a band at 1335 cm-1, that corresponds to C-O

angular deformations of phenols, at 2830 and 2921 cm−<sup>1</sup> due to symmetric and asymmetric C-H vibration respectively and 3269 cm−<sup>1</sup> from O-H stretching vibration. In BCS samples there are two additional peaks, at 1710 cm−<sup>1</sup> from a C=O stretching vibration and a 1180 cm−<sup>1</sup> that correspond to C-O symmetric vibration, this is indicative that there are other acyl compounds in the extract that are account for a major proportion on the surface. *Molecules* **2018**, *23*, x FOR PEER REVIEW 3 of 16 O symmetric vibration, this is indicative that there are other acyl compounds in the extract that are account for a major proportion on the surface.

Becke 3-Lee-Yang-Parr (B3LYP) model simulation results are shown in Figure 1b, considering the cyanidin 3-O-glucoside molecule. The bands obtained are 3370 and 3290 cm−<sup>1</sup> (O-H symmetric stretching vibration), 3220 cm−<sup>1</sup> (C-H symmetric stretching), 1719 and 1689 cm−<sup>1</sup> (C=C scissoring on pyran and phenolic group respectively), 1535 cm−<sup>1</sup> (C-H scissoring), 1419 cm−<sup>1</sup> (asymmetric ring vibration on plane), 1380 cm−<sup>1</sup> (C-H deformation), 1213 cm−<sup>1</sup> (C-O stretching), 1154 cm−<sup>1</sup> (a scissoring plane vibration of phenol ring), 959 cm−<sup>1</sup> (phenol ring C-H asymmetric stretching), 861 cm−<sup>1</sup> (C-H phenol ring symmetric stretching) and 738 cm−<sup>1</sup> (phenol ring C-H deformation). Becke 3-Lee-Yang-Parr (B3LYP) model simulation results are shown in Figure 1b, considering the cyanidin 3-O-glucoside molecule. The bands obtained are 3370 and 3290 cm−1 (O-H symmetric stretching vibration), 3220 cm−1 (C-H symmetric stretching), 1719 and 1689 cm−1 (C=C scissoring on pyran and phenolic group respectively), 1535 cm−1 (C-H scissoring), 1419 cm−1 (asymmetric ring vibration on plane), 1380 cm−1 (C-H deformation), 1213 cm−1 (C-O stretching), 1154 cm−1 (a scissoring plane vibration of phenol ring), 959 cm−1 (phenol ring C-H asymmetric stretching), 861 cm−1 (C-H phenol ring symmetric stretching) and 738 cm−1 (phenol ring C-H deformation).

**Figure 1.** (**a**) Experimental FTIR spectra of black carrot extracts; samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), (**b**) Simulated FTIR spectra using B3LYP calculation and cyanidin 3-O-glucoside molecule (used for simulation). **Figure 1.** (**a**) Experimental FTIR spectra of black carrot extracts; samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), (**b**) Simulated FTIR spectra using B3LYP calculation and cyanidin 3-O-glucoside molecule (used for simulation).

These results suggest that cyanidin 3-O-glucoside is the major anthocyanin component in black carrot, since they share similar vibrational frequencies and the same functional groups, like previously reported values for black carrot [20–23]. To confirm this notion, high performance liquid chromatography (HPLC) was performed for samples BCS and BCA. Figure 2a shows the HPLC chromatogram from sample BCS and Figure 2b shows the HPLC chromatogram for BCA. Peak These results suggest that cyanidin 3-O-glucoside is the major anthocyanin component in black carrot, since they share similar vibrational frequencies and the same functional groups, like previously reported values for black carrot [20–23]. To confirm this notion, high performance liquid chromatography (HPLC) was performed for samples BCS and BCA. Figure 2a shows the HPLC chromatogram from sample BCS and Figure 2b shows the HPLC chromatogram for BCA. Peak

identification was made using previous reports [22–24]. Table 1 shows the retention times of the

characteristic peaks and compounds identified.

identification was made using previous reports [22–24]. Table 1 shows the retention times of the characteristic peaks and compounds identified. *Molecules* **2018**, *23*, x FOR PEER REVIEW 4 of 16

**Figure 2.** High performance liquid chromatography (HPLC) of black carrot extracts: (**a**) sample BCS (extracted with ethanol/citric acid). (**b**) sample BCA (extracted with ethanol/acetic acid). Peaks were identified and are shown in Table 1. **Figure 2.** High performance liquid chromatography (HPLC) of black carrot extracts: (**a**) sample BCS (extracted with ethanol/citric acid). (**b**) sample BCA (extracted with ethanol/acetic acid). Peaks were identified and are shown in Table 1.

The chromatogram of sample BCS shows two additional peaks (anthocyanins reported as derived from feluric acid) compared with sample BCA. HPLC analysis confirmed the presence of cyanidin 3-O-glucoside molecule and its derivatives in the black carrot extracts obtained. The chromatogram of sample BCS shows two additional peaks (anthocyanins reported as derived from feluric acid) compared with sample BCA. HPLC analysis confirmed the presence of cyanidin 3-O-glucoside molecule and its derivatives in the black carrot extracts obtained.


**Table 1.** Retention time and anthocyanin identification from black carrot extracts BCS (extracted with **Table 1.** Retention time and anthocyanin identification from black carrot extracts BCS (extracted with ethanol/citric acid) and BCA (extracted with ethanol/acetic acid).

7 - 25.17 Ferulic acid derivative of peonidin 3-xylosyl-glucosyl-galactoside Figure 3 shows the total anthocyanin content obtained by a differential pH analytical method. Using ethanol with acetic acid as extraction solvent (sample BCA) leads to more anthocyanin content compared with using ethanol and citric acid, which is congruent with results published before on anthocyanin quantification of extracts from other plants [24,25]. Anthocyanin content is reduced significantly after passing through an Amberlite XAD7 resin column (Figure 3a); this could be Figure 3 shows the total anthocyanin content obtained by a differential pH analytical method. Using ethanol with acetic acid as extraction solvent (sample BCA) leads to more anthocyanin content compared with using ethanol and citric acid, which is congruent with results published before on anthocyanin quantification of extracts from other plants [24,25]. Anthocyanin content is reduced significantly after passing through an Amberlite XAD7 resin column (Figure 3a); this could be explained as a natural degradation process under the experimental conditions. Further chromatographic studies should investigate if 3-O-glucoside anthocyanin derivatives could be trapped in the resin.

6 - 23.98 Feruic acid derivative of pelargonidin 3-xylosyl-glucosyl-galactoside

explained as a natural degradation process under the experimental conditions. Further chromatographic studies should investigate if 3-O-glucoside anthocyanin derivatives could be trapped in the resin. The four extracts (BCA, BCS, BCAR, and BCSR) were microencapsulated using TEOS and their anthocyanin content is shown in Figure 3b; there was an average loss of 7 ± 0.1% and 2.5 ± 0.005% in the anthocyanin content for samples without passing through the resin column (TBCS, TBCA) and samples after passing through the resin column (TBCSR, TBCAR), respectively. That suggests that TEOS incorporation has a negligible effect in preventing degradation. This could be explained by hydrolysis and condensation processes of the alkoxide that could be favored by functional groups of the acids used. Also, bonds between Si-O and 3-O-glucoside structure, as proposed by other authors [1,3], could lead to less total anthocyanin content in microencapsulated samples than in samples without microencapsulation.

Microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR).

identified and are shown in Table 1.

*Molecules* **2018**, *23*, x FOR PEER REVIEW 4 of 16

**Figure 2.** High performance liquid chromatography (HPLC) of black carrot extracts: (**a**) sample BCS (extracted with ethanol/citric acid). (**b**) sample BCA (extracted with ethanol/acetic acid). Peaks were

**Table 1.** Retention time and anthocyanin identification from black carrot extracts BCS (extracted with

3 16.91 16.88 Sinapic acid derivative of cyanidin 3-xylosyl-glucosyl-galactoside 4 19.43 19.42 Ferulic acid derivative of cyanidin 3-xylosyl-glucosyl-galactoside 5 20.45 20.44 Coumaric acid derivative of cyanidin 3-xylosyl-glucosyl-galactoside 6 - 23.98 Feruic acid derivative of pelargonidin 3-xylosyl-glucosyl-galactoside 7 - 25.17 Ferulic acid derivative of peonidin 3-xylosyl-glucosyl-galactoside

Figure 3 shows the total anthocyanin content obtained by a differential pH analytical method. Using ethanol with acetic acid as extraction solvent (sample BCA) leads to more anthocyanin content compared with using ethanol and citric acid, which is congruent with results published before on anthocyanin quantification of extracts from other plants [24,25]. Anthocyanin content is reduced significantly after passing through an Amberlite XAD7 resin column (Figure 3a); this could be explained as a natural degradation process under the experimental conditions. Further

cyanidin 3-O-glucoside molecule and its derivatives in the black carrot extracts obtained.

1 10.94 10.93 Cyanidin-3-xylosyl-glucosyl-galactoside 2 14.46 14.39 Cyanidin-3-xylosyl-galactoside

ethanol/citric acid) and BCA (extracted with ethanol/acetic acid).

**Peak Retention Time (min) Anthocyanin BCS BCA** 

The chromatogram of sample BCS shows two additional peaks (anthocyanins reported as derived from feluric acid) compared with sample BCA. HPLC analysis confirmed the presence of

**Figure 3.** Anthocyanin content in black carrot extracts. (**a**) Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), and samples after resin column (BCSR and BCAR). (**b**) Microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR). **Figure 3.** Anthocyanin content in black carrot extracts. (**a**) Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), and samples after resin column (BCSR and BCAR). (**b**) Microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR). hydrolysis and condensation processes of the alkoxide that could be favored by functional groups of the acids used. Also, bonds between Si-O and 3-O-glucoside structure, as proposed by other authors [1,3], could lead to less total anthocyanin content in microencapsulated samples than in samples without microencapsulation.

The anthocyanin contents for BCS, BCSR and BCAR and the same microencapsulated samples are in accordance with other microencapsulated powder samples reported for black carrot extracts [8]. BCA and TBCA extracts had the highest anthocyanin content compared with previously mentioned extracts, but it was not as high as the extraction reported using enzymes for other subspecies of black carrot [12,18], nevertheless the extraction method reported here, is inexpensive compared with others and it could be competitive for several industrial applications. The anthocyanin contents for BCS, BCSR and BCAR and the same microencapsulated samples are in accordance with other microencapsulated powder samples reported for black carrot extracts [8]. BCA and TBCA extracts had the highest anthocyanin content compared with previously mentioned extracts, but it was not as high as the extraction reported using enzymes for other subspecies of black carrot [12,18], nevertheless the extraction method reported here, is inexpensive compared with others and it could be competitive for several industrial applications.

#### *2.2. UV Radiation Study 2.2. UV Radiation Study*

*2.3. Thermal Stability* 

In the UV radiation stability test (Figure 4a), there was a reduction of 19.81% and 17.99% of the total anthocyanin content for BCS and BCA, respectively. On the other hand, the extracts under resin purification BCSR and BCAR had a reduction of 7.06% and 12.55% from its total anthocyanin content. Considering statistical variations, it cannot be ensured that anthocyanin acylation results in protection against UV radiation. In the UV radiation stability test (Figure 4a), there was a reduction of 19.81% and 17.99% of the total anthocyanin content for BCS and BCA, respectively. On the other hand, the extracts under resin purification BCSR and BCAR had a reduction of 7.06% and 12.55% from its total anthocyanin content. Considering statistical variations, it cannot be ensured that anthocyanin acylation results in protection against UV radiation.

**Figure 4.** Anthocyanin content in black carrot extracts as function of time under UV radiation (315– 400 nm); (**a**) Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), and samples after resin column (BCSR and BCAR). (**b**) Microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR). **Figure 4.** Anthocyanin content in black carrot extracts as function of time under UV radiation (315–400 nm); (**a**) Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), and samples after resin column (BCSR and BCAR). (**b**) Microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR).

Figure 4b shows samples microencapsulated with TEOS, and they follow a quasi-linear decay behavior instead an exponential decay (non-microencapsulated extracts). This can be seen in the reduction of total anthocyanin content which was 21.63%, 20.17%, 22.68%, and 22.82% for TBCS, TBCA, TBCSR and TBCAR, respectively, that are higher losses compared with samples without microencapsulation. Figure 4b shows samples microencapsulated with TEOS, and they follow a quasi-linear decay behavior instead an exponential decay (non-microencapsulated extracts). This can be seen in the reduction of total anthocyanin content which was 21.63%, 20.17%, 22.68%, and 22.82% for TBCS, TBCA, TBCSR and TBCAR, respectively, that are higher losses compared with samples without microencapsulation.

ethanol/citric acid as extraction solvent; the first ones have a greater decay rate because their graphs (Figure 5) have a greater slope. This slope difference is more visible at high temperatures. On the other hand, considering anthocyanin content loss percentages of 9.28% and 9.31% at 40 °C, 20.08%

Sample BCA presents same trend compared with BCAR in thermal stability at different

#### *2.3. Thermal Stability*

Sample BCA presents same trend compared with BCAR in thermal stability at different temperatures (Figure 5a–c). The same behavior was obtained for BCS and BCSR, therefore purifying extracts through resin column does not have a significant improvement in thermal stability. There is a difference in the behavior of the decay between extracts obtained with ethanol/acetic acid and ethanol/citric acid as extraction solvent; the first ones have a greater decay rate because their graphs (Figure 5) have a greater slope. This slope difference is more visible at high temperatures. On the other hand, considering anthocyanin content loss percentages of 9.28% and 9.31% at 40 ◦C, 20.08% and 23.96% at 60 ◦C, and 29.53% and 47.07% at 80 ◦C for BCS and BCA, respectively. The thermal stability behavior of anthocyanins as a function of time at 40 and 60 ◦C for TBCA and TBCAR samples is similar (Figure 5d,e). TBCS and TBCSR also had the same trend in thermal stability curves at 40 and 60 ◦C, from this behavior we assume that the resin column did not influence the thermal stability. Nevertheless, at 80 ◦C ethanolic extraction with acetic acid and encapsulated samples showed a rapid decay in anthocyanin content. Comparing the total anthocyanins loss of the encapsulated and non-encapsulated samples (9.28% and 14.93% for BCS and TBCS; 9.31% and 8.2% for BCA and TBCA, respectively) at 40 ◦C, it is possible to conclude that microencapsulation does not prevent thermal degradation in this case. In higher temperatures the same conclusion was obtained (for example at 80 ◦C: 29.53% and 35.91% for BCS and TBCS, 45.07% and 57.15% for BCA and TBCA, respectively). *Molecules* **2018**, *23*, x FOR PEER REVIEW 6 of 16 and 23.96% at 60 °C, and 29.53% and 47.07% at 80 °C for BCS and BCA, respectively. The thermal stability behavior of anthocyanins as a function of time at 40 and 60 °C for TBCA and TBCAR samples is similar (Figures 5d,e). TBCS and TBCSR also had the same trend in thermal stability curves at 40 and 60 °C, from this behavior we assume that the resin column did not influence the thermal stability. Nevertheless, at 80 °C ethanolic extraction with acetic acid and encapsulated samples showed a rapid decay in anthocyanin content. Comparing the total anthocyanins loss of the encapsulated and nonencapsulated samples (9.28% and 14.93% for BCS and TBCS; 9.31% and 8.2% for BCA and TBCA, respectively) at 40 °C, it is possible to conclude that microencapsulation does not prevent thermal degradation in this case. In higher temperatures the same conclusion was obtained (for example at 80 °C: 29.53% and 35.91% for BCS and TBCS, 45.07% and 57.15% for BCA and TBCA, respectively).

**Figure 5.** Anthocyanin content as function of time at different temperatures; (**a**,**d**) at 40 °C, (**b**,**e**) at 60 °C, and (**c**,**f**) at 80 °C. Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), samples after resin column (BCSR and BCAR), and microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR). **Figure 5.** Anthocyanin content as function of time at different temperatures; (**a**,**d**) at 40 ◦C, (**b**,**e**) at 60 ◦C, and (**c**,**f**) at 80 ◦C. Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), samples after resin column (BCSR and BCAR), and microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR).

These results showed that the anthocyanins were more stable using ethanol with citric acid (as extraction solvent) and purified with resin at 40 °C (BCSR) and the stability can be slightly enhanced when TEOS microencapsulation is used at high temperatures. In this experiment, BCAR (extraction of ethanol acidified with acetic acid after purification) had the highest value of degradation and it is

These results showed that the anthocyanins were more stable using ethanol with citric acid (as extraction solvent) and purified with resin at 40 ◦C (BCSR) and the stability can be slightly enhanced when TEOS microencapsulation is used at high temperatures. In this experiment, BCAR (extraction of ethanol acidified with acetic acid after purification) had the highest value of degradation and it is more clearly when microencapsulation with TEOS is used. For samples where resin is used, the results are consistent, even using TEOS for microencapsulation. *Molecules* **2018**, *23*, x FOR PEER REVIEW 7 of 16 more clearly when microencapsulation with TEOS is used. For samples where resin is used, the results are consistent, even using TEOS for microencapsulation.

#### *2.4. pH Storage Stability 2.4. pH Storage Stability*

The pH changes were evaluated using a short-term storage test during five days for analyzing monomeric anthocyanin content changes at acidic, neutral and alkaline pH and Figure 6 shows these changes. The pH changes were evaluated using a short-term storage test during five days for analyzing monomeric anthocyanin content changes at acidic, neutral and alkaline pH and Figure 6 shows these changes.

**Figure 6.** Anthocyanin content as function of storage time at different pH buffers; (**a**,**d**) pH = 4, (**b**,**e**) pH = 7 and (**c**,**f**) pH = 10. Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), samples after resin column (BCSR and BCAR), and microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR). **Figure 6.** Anthocyanin content as function of storage time at different pH buffers; (**a**,**d**) pH = 4, (**b**,**e**) pH = 7 and (**c**,**f**) pH = 10. Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), samples after resin column (BCSR and BCAR), and microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR).

and BCAR 3.17%, and the major changes for the extracts were at alkaline pH (pH = 10, Figure 6c), since the samples with a higher loss from its initial value were 83.38% and 26.98% for BCS and BCA;

At acid pH (pH = 4, Figure 6a) there was an anthocyanin content loss of 4.98% and 3.22% for BCS and BCA, respectively, and for resin purified extracts (BCSR and BCAR) the losses were 5.58% and

At acid pH (pH = 4, Figure 6a) there was an anthocyanin content loss of 4.98% and 3.22% for BCS and BCA, respectively, and for resin purified extracts (BCSR and BCAR) the losses were 5.58% and 3.01%. At neutral pH (pH = 7, Figure 6b), the content loss was BCS 13.76%, BCA 4.39%, BCSR 5.60% and BCAR 3.17%, and the major changes for the extracts were at alkaline pH (pH = 10, Figure 6c), since the samples with a higher loss from its initial value were 83.38% and 26.98% for BCS and BCA; at this pH, purified extracts had less degradation with a content loss of 15.57% for BCSR and 5.38% BCAR.

For microencapsulated samples at acid pH (Figure 6d) there was an anthocyanin content loss of 1.69% and 2.63% for TBCS and TBCA respectively and for TBCSR and TBCAR the loss was 5.36% and 3.10%. At neutral pH (Figure 6e) the content loss was 18.02% for TBCS, 7.8% for TBCAR, 7.7% for TBCSR and 8.39% for TBCAR and finally at alkaline pH (Figure 6f) major changes in content were found, such as 86.07% for TBCS, 33.48% for TBCA, 19.90% for TBCSR and 15.02% for TBCAR. Therefore, degradation of microencapsulated samples was reduced at acid pH. When the pH increases, degradation increases too due to the increase in alkalinity.

In both cases, microencapsulated and non-microencapsulated samples, the graphical tendency is the same and anthocyanins in BCAR samples were the most stable after the elapsed time under three-different conditions of pH, indicating that the anthocyanins extracted with ethanol/acetic acid are more stable to pH changes in comparison with the ethanol/citric acid extracts (higher anthocyanin content loss), which is highly evident at pH = 10.

Since the results from degradation studies of samples microencapsulated with TEOS showed no significant improvement in the stability of the extracts (except for the experiment at 60 ◦C), these samples were not analyzed for antioxidant activity and color in the food models. This decision was made considering also that the activity of nutraceutical compounds was reduced after treatment [3].

#### *2.5. DPPH and Electrochemical CUPRAC Antioxidant Content Test*

As seen from Figure 7a there is a direct relationship of anthocyanin content and antiradical activity; when the anthocyanin content is higher, the antioxidant effect increases. With more phenolic compounds, such as anthocyanins, a higher antiradical activity is expected.

The extract obtained with acetic acid leads to a higher yield of antiradical activity (614.52 µM TE g fw−<sup>1</sup> ) which is 15.5% higher compared with the extract obtained with citric acid. This was expected because the acylated nature of the extracted anthocyanin [25] confers higher antiradical activity than monomeric anthocyanins; also, the use of XAD7 resin reduced antiradical activity on 20.20% and 18.94% for BCS and BCA extracts, respectively.

In the case of CUPRAC test, Figure 7b shows cyclic voltammograms for the antioxidant agents obtained with different extraction methods. It can be observed that the initial potential for the BCAR and BCA was 0.433 and 0.428 V, respectively. Those values exhibited a shift to negative potentials compared with 0.454 and 0.488 V of BCSR and BCS, respectively. This shift to potential negative values is related with an increment in the amount of the complex *Cu*(*Nc*) + 2 due to the capability of the antioxidant agent to donate an electron to the oxidized complex *Cu*(*Nc*) 2+ 2 according to the following equation:

$$a\mathbb{C}u(\mathrm{Nc})\_2^{2+} + bAO\_{\mathrm{red}} \leftrightarrow c\mathbb{C}u(\mathrm{Nc})\_2^+ + dAO\_{\mathrm{ox}}^+ \tag{1}$$

where the *AOred* is the reduced antioxidant agent and the *AO*<sup>+</sup> *oxi* is the antioxidant agent when it was oxidized. This behavior is directly related with the antioxidant agent capability of the samples to promote the reduction reaction for the molecule that was previously oxidized (*Cu*(*Nc*) 2+ 2 )*.* Also, the description above was based on the Nernst equation:

$$E = \, ^0E + \frac{RT}{F} \ln \frac{a\_{Cu(Nc)\_2^{2+}}}{a\_{Cu(Nc)\_2^{+}}} \tag{2}$$

where the potential of the reaction on the equilibrium were shifted to negative values owing an increase in the activity (concentration of *Cu*(*Nc*) 2+ 2 ) of the products.

BCAR.

*Molecules* **2018**, *23*, x FOR PEER REVIEW 8 of 16

at this pH, purified extracts had less degradation with a content loss of 15.57% for BCSR and 5.38%

increases, degradation increases too due to the increase in alkalinity.

*2.5. DPPH and Electrochemical CUPRAC Antioxidant Content Test* 

content loss), which is highly evident at pH = 10.

For microencapsulated samples at acid pH (Figure 6d) there was an anthocyanin content loss of 1.69% and 2.63% for TBCS and TBCA respectively and for TBCSR and TBCAR the loss was 5.36% and 3.10%. At neutral pH (Figure 6e) the content loss was 18.02% for TBCS, 7.8% for TBCAR, 7.7% for TBCSR and 8.39% for TBCAR and finally at alkaline pH (Figure 6f) major changes in content were found, such as 86.07% for TBCS, 33.48% for TBCA, 19.90% for TBCSR and 15.02% for TBCAR. Therefore, degradation of microencapsulated samples was reduced at acid pH. When the pH

In both cases, microencapsulated and non-microencapsulated samples, the graphical tendency is the same and anthocyanins in BCAR samples were the most stable after the elapsed time under three-different conditions of pH, indicating that the anthocyanins extracted with ethanol/acetic acid are more stable to pH changes in comparison with the ethanol/citric acid extracts (higher anthocyanin

Since the results from degradation studies of samples microencapsulated with TEOS showed no significant improvement in the stability of the extracts (except for the experiment at 60 °C), these samples were not analyzed for antioxidant activity and color in the food models. This decision was made considering also that the activity of nutraceutical compounds was reduced after treatment [3].

As seen from Figure 7a there is a direct relationship of anthocyanin content and antiradical activity; when the anthocyanin content is higher, the antioxidant effect increases. With more phenolic

**Figure 7.** (**a**) Antiradical activity of black carrot extracts using DPPH assay at different concentrations. (**b**) Cyclic voltammograms for the four extracts (400 µM) in a CUPRAC solution. The scan began at the open circuit potential (OCP) with a sweep velocity of 100 mV·s−1. (**c**) Electrochemical antioxidant activity and compared with Trolox activity at the same concentration. Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), samples after resin column (BCSR and BCAR). **Figure 7.** (**a**) Antiradical activity of black carrot extracts using DPPH assay at different concentrations. (**b**) Cyclic voltammograms for the four extracts (400 µM) in a CUPRAC solution. The scan began at the open circuit potential (OCP) with a sweep velocity of 100 mV·s −1 . (**c**) Electrochemical antioxidant activity and compared with Trolox activity at the same concentration. Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), samples after resin column (BCSR and BCAR).

Furthermore, the peak current for the oxidation process in the voltammograms has a direct relation with the concentration of *Cu*(*Nc*) + 2 because if the electrolyte has a higher amount of *Cu*(*Nc*) + 2 we obtained a major amount of oxidizer molecules (*Cu*(*Nc*) 2+ 2 ) resulting in an increment of the current value. This is possible because the current is directly proportional for the concentration of the species in the reaction (*i* ∝ *C*) [26,27].

The Trolox calibration curve was obtained from the oxidation peak current and was used to analyze the antioxidant activity for the black carrot extracts obtained by the different extraction methods (*r* <sup>2</sup> = 0.9768). It is proposed that ratio between current peak of the black carrot and the current peak of Trolox (at same concentration, 400 µM) allows determining the antioxidant capability of each black carrot extract. This method is similar to calculating the Trolox Equivalent Antioxidant Capacity (TEAC), where calculations were made by the ratio of molar absorptivity of problem species and Trolox under the corresponding conditions [28]. In order to have cyanidin 3-O-glucoside at a 400 µM concentration, samples were diluted (Figure 3a) and the molecular weight of the anthocyanin was used for the analytical pH differential method.

The calculated Electrochemical Antioxidant Capacity (EAC) is shown in Figure 7c, giving the following results: 0.684, 0.346, 0.748 and 0.591 for BCS, BCA, BCSR and BCAR, respectively. These EAC results exhibited a behavior in accordance with the DPPH colorimetric method. BCA extract had the highest antiradical activity of all samples, followed by BCS, then BCAR and finally BCSR. Also, analyzing the antioxidant activity for black carrot extracts, values of current were under the values of the Trolox calibration plot. The decrease of the antiradical activity is not as higher than the anthocyanin content in Figure 3a; this is because electrochemical CUPRAC methods measure compounds related with the antiradical activity at an electron level. These CUPRAC test shows that our samples could

color.

except BCA.

25 days in storage.

have several 3,7-diglucoside derivatives and other phenolic compounds. The antiradical activity of black carrots extracts was higher than other reported values of several extracts [29]. *Molecules* **2018**, *23*, x FOR PEER REVIEW 10 of 16

#### *2.6. Color of Black Carrot Extracts on Food Models* brown color tendency (hue angle below 2°) at the beginning of the experiment. The jelly has more

Figure 8a−d show the average results from the image analysis for food models using the black carrot extracts and Red FD&C analysis for comparison. All samples (in yogurt and jelly), had a light brown color tendency (hue angle below 2◦ ) at the beginning of the experiment. The jelly has more saturated colors than yogurt and this is due to the base color of the food models (white vs. pale yellow). Red FD&C had the highest luminosity in yogurt and is more saturated than BCS and BCA samples (chroma value). FD&C in jelly is darker, but has the same color saturation than BCS, i.e. in the food model; FD&C and BCS have the same color saturation for the human eye. BCA sample is less saturated in jelly. In the case of BCSR and BCAR, in yogurt, the color is less saturated but darker; and for jelly, they have almost the same saturation (slightly less saturated) but it has a much brighter color. saturated colors than yogurt and this is due to the base color of the food models (white vs. pale yellow). Red FD&C had the highest luminosity in yogurt and is more saturated than BCS and BCA samples (chroma value). FD&C in jelly is darker, but has the same color saturation than BCS, i.e. in the food model; FD&C and BCS have the same color saturation for the human eye. BCA sample is less saturated in jelly. In the case of BCSR and BCAR, in yogurt, the color is less saturated but darker; and for jelly, they have almost the same saturation (slightly less saturated) but it has a much brighter

**Figure 8.** (**a**,**c**) Sectional polar and (**b**,**d**) cartesian diagram of color for the black carrots extracts and the red FD&C on food models (yogurt and jelly) at 0 and 25 days of storage. **Figure 8.** (**a**,**c**) Sectional polar and (**b**,**d**) cartesian diagram of color for the black carrots extracts and the red FD&C on food models (yogurt and jelly) at 0 and 25 days of storage.

As seen in Table 2, after the 25-day storage time the color differences (ΔE) in yogurt, of all samples, have values higher than five, which indicates that the color difference at the beginning and after the elapsed time is visually evident; also, samples have higher saturation (lowest chroma value) As seen in Table 2, after the 25-day storage time the color differences (∆E) in yogurt, of all samples, have values higher than five, which indicates that the color difference at the beginning and after the elapsed time is visually evident; also, samples have higher saturation (lowest chroma value) after storage and specifically the black carrots extract samples get darker since L\* value is lower. BCS samples showed similar color difference after storage time compared with red FD&C.

after storage and specifically the black carrots extract samples get darker since L\* value is lower. BCS

difference could be distinguished but was not as evident as the rest of the samples; also, BCA samples are the only ones with different saturation and luminosity trends, they had lower chroma values and

For jelly samples, only sample BCA has a value lower than five, which indicates that the color

samples showed similar color difference after storage time compared with red FD&C.

**Table 2.** Color comparison between food models (yogurt and jelly) using black carrot extracts after

**Sample ΔE Yogurt ΔE Jelly**  Red FD&C 12.62 10.92 BCS 16.45 17.61 BCSR 24.27 9.21 BCA 18.31 2.82 BCAR 25.12 12.71

For yogurt, BCSR and BCAR samples have the highest color change indicating that cyanidin-3-

glucoside derivative anthocyanins are not suitable to be used in this food model, because the acetate


**Table 2.** Color comparison between food models (yogurt and jelly) using black carrot extracts after 25 days in storage.

For jelly samples, only sample BCA has a value lower than five, which indicates that the color difference could be distinguished but was not as evident as the rest of the samples; also, BCA samples are the only ones with different saturation and luminosity trends, they had lower chroma values and are darker. The rest of the samples are less saturated (higher chroma value) and brighter. The samples with resin purification (BCSR and BCAR) had a similar color difference compared to red FD&C, except BCA.

For yogurt, BCSR and BCAR samples have the highest color change indicating that cyanidin-3-glucoside derivative anthocyanins are not suitable to be used in this food model, because the acetate group caused important appearance differences under these conditions; for jelly, BCS and BCAR had the highest color changes but BCSR and BCA the lowest compared with Red FD&C. Pigment concentration of the samples must change for the specific commercial use, for instance, strawberry yogurt samples have different L\*, a\* and b\* values [3].

#### **3. Materials and Methods**

Fresh black carrot (*Daucus Carota* var. *L.* ssp. *sativus* var. *atrorubens* Alef.) were cultivated in Tlaxcala, Mexico and donated by a local cultivator, then were washed with tap water and stored in 3 kg perforated plastics bags and kept at −20 ◦C until further use. All reactants used were analytical grade. Ethanol 99.5%, citric acid 99.5%, Amberlite XAD7 resin, tetraethyl orthosilicate (TEOS) 98%, potassium chloride, 6-hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox), and 2,2-diphenyl-1-picrylhydrazyl (DPPH), CuCl2·2H2O (99%), neocuproine, and ammonium acetate (98.0%) were supplied by Sigma Aldrich (Toluca, Mexico). Acetic acid, sodium acetate, hydrochloric acid, buffer solutions (pH: 4, 7 & 10) and potassium hydroxide were supplied by J.T. Baker (Mexico City, Mexico). Methanol was supplied by Macron (Hamilton, PA, USA). All experiments were carried out using distilled water obtained from an Elix Advantage Water Purification System (Queretaro, Mexico).

#### *3.1. Anthocyanins Extraction and TEOS Microencapsulation*

Acidified ethanol was used as extraction solvent for all samples. Anthocyanins were obtained by blending 150 mL of acidified ethanol (citric/acetic acid solution 85:15 *v*/*v*) and 150 g of sliced frozen carrots (without thawing) with a Grinder 6807 blender Oster (Mexico) for 20 min. Solids were removed by filtration using a 100-mesh sieve filter. The liquid phase (extract) was labeled as BCS (acidified with citric acid) and BCA (acidified with acetic acid). Extracts BCS and BCA were introduced, separately, to a resin column containing Amberlite XAD7 resin for removing non-aromatic compounds from the extract. Flow rate was 32 mL/min (20BV/h) and a solution of 95 mL of ethanol and 5 mL of acidic water (pH = 1.4) was used as eluent. Finally, solvent excess was evaporated at 40 ◦C using vacuum (R-100, Büchi, Mexico). Samples obtained after this procedure were labeled BCSR and BCAR. All samples were stored in sealed amber glass vials at 4 ◦C until further use. Then samples were submitted to microencapsulation using TEOS with a procedure reported elsewhere [1,3]. The samples after microencapsulation were labeled as TBCS, TBCA, TBCSR and TBCAR. A flow diagram is shown in Figure 9.

*Molecules* **2018**, *23*, x FOR PEER REVIEW 11 of 16

group caused important appearance differences under these conditions; for jelly, BCS and BCAR had the highest color changes but BCSR and BCA the lowest compared with Red FD&C. Pigment concentration of the samples must change for the specific commercial use, for instance, strawberry

Fresh black carrot (*Daucus Carota* var. *L.* ssp. *sativus* var. *atrorubens* Alef.) were cultivated in

Acidified ethanol was used as extraction solvent for all samples. Anthocyanins were obtained

by blending 150 mL of acidified ethanol (citric/acetic acid solution 85:15 *v*/*v*) and 150 g of sliced frozen carrots (without thawing) with a Grinder 6807 blender Oster (Mexico) for 20 min. Solids were removed by filtration using a 100-mesh sieve filter. The liquid phase (extract) was labeled as BCS (acidified with citric acid) and BCA (acidified with acetic acid). Extracts BCS and BCA were introduced, separately, to a resin column containing Amberlite XAD7 resin for removing nonaromatic compounds from the extract. Flow rate was 32 mL/min (20BV/h) and a solution of 95 mL of ethanol and 5 mL of acidic water (pH = 1.4) was used as eluent. Finally, solvent excess was evaporated at 40 °C using vacuum (R-100, Büchi, Mexico). Samples obtained after this procedure were labeled BCSR and BCAR. All samples were stored in sealed amber glass vials at 4 °C until further use. Then samples were submitted to microencapsulation using TEOS with a procedure reported elsewhere

Tlaxcala, Mexico and donated by a local cultivator, then were washed with tap water and stored in 3 kg perforated plastics bags and kept at −20 °C until further use. All reactants used were analytical grade. Ethanol 99.5%, citric acid 99.5%, Amberlite XAD7 resin, tetraethyl orthosilicate (TEOS) 98%, potassium chloride, 6-hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox), and 2,2 diphenyl-1-picrylhydrazyl (DPPH), CuCl2·2H2O (99%), neocuproine, and ammonium acetate (98.0%) were supplied by Sigma Aldrich (Toluca, Mexico). Acetic acid, sodium acetate, hydrochloric acid, buffer solutions (pH: 4, 7 & 10) and potassium hydroxide were supplied by J.T. Baker (Mexico City, Mexico). Methanol was supplied by Macron (Hamilton, PA, USA). All experiments were carried out using distilled water obtained from an Elix Advantage Water Purification System (Queretaro,

yogurt samples have different L\*, a\* and b\* values [3].

*3.1. Anthocyanins Extraction and TEOS Microencapsulation* 

**3. Materials and Methods** 

Mexico).

**Figure 9.** Flow diagram of experimental processes for obtaining samples. Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), samples after resin column (BCSR and BCAR), and microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR). **Figure 9.** Flow diagram of experimental processes for obtaining samples. Samples extracted with ethanol/citric acid (BCS) and ethanol/acetic acid (BCA), samples after resin column (BCSR and BCAR), and microencapsulated samples using TEOS (TBCS, TBCA, TBCSR, TBCAR).

#### *3.2. Anthocyanin Content*

It has been reported that black carrot contains different monomeric anthocyanins with different sugar moieties such as peonidin, pelargonidin and cyanidin, being cyanidin 3-O-R the major compound in the total anthocyanin content [20–23]. In order to confirm this and to justify the use of cyanidin 3-O-glucoside for analytical measurements, Fourier Transform Infrared Spectroscopy (FTIR) spectra under attenuated total reflectance (ATR) was measured by a Frontier MIR/NIR Spectrometer (Perkin Elmer, Waltham, MA, United States), for dried samples of BCA and BCS and then compared with a frequency quantum chemical calculation of the same molecule (Figure 1c).

For the frequency calculation of cyanidin 3-O-glucoside molecule a personal computer running Gaussian 98W [30] was used. The geometry was fully optimized assuming Cs point group symmetry using the Becke 3-Lee-Yang-Parr (B3LYP), supplemented with the standard 6-31 + G basis sets. The simulated IR spectra were plotted using Avogadro molecular viewer [31] and the vibrational modes were analyzed and compared to the experimental data as mentioned.

Total anthocyanins content (monomeric anthocyanins) was determined using the pH-differential method reported by Giusti and Wrolstad [32], using a molar extinction coefficient of 26900 M-1cm-1 that corresponds to cyanidin 3-O-glucoside. The average molecular weight used was 756.87 g mol-1 of the anthocyanins according to previous studies from black carrot [32–34]. A VWR 1600-PC spectrophotometer and 1 cm path length glass cells were used; measurements were performed scanning from 700 to 400 nm at room temperature (~24 ◦C). Finally, High pressure liquid chromatography (HPLC) was performed on a Flexar LC (Perkin Elmer) system using a 250 mm × 4.6 mm C18 reverse phase column and BCS and BCA samples were measured according to conditions reported elsewhere [34,35]; using a flow rate of 1.0 mL/min and the chromatographs were recorded at 520 nm using PDA Plus Detector coupled with the equipment.

#### *3.3. Degradation Studies*

Black carrot extracts were transferred into vials with screw caps to perform degradation tests. All experiments were performed in triplicate and the referred anthocyanin content was normalized to 200 mg/L in all the samples in order to make direct comparison between samples so the results are presented as remaining percentage of cyanidin 3-O-glycoside.

The ASTM D 4320 method was used to determine anthocyanin stability against UV radiation. Black carrot extracts were exposed to UV lamp irradiation (315–400 nm) for 160 min; samples were placed within 15 cm from the source. Measurements were taken every 40 minand the temperature was kept constant at 25 ◦C under a working area of 3.5 m<sup>2</sup> isolated from other light sources [1,3].

For thermal stability studies, samples were placed in a preheated water bath at 80 ◦C, 60 ◦C and 40 ◦C. Samples were removed from water bath every 20 min, up to 120 minand rapidly cooled to room temperature. Anthocyanin content was analyzed immediately [35].

The anthocyanin stability in storage was also studied at three different pH conditions (4, 7 and 10) at room temperature (~24 ◦C). For this purpose, 3 mL of phosphate buffer solution were prepared at the required pH conditions and then colored with 500 µL of black carrot extract concentrate; these colored solutions were used without further treatment. Finally, anthocyanin storage stability was determined every 24 h during 5 days for each of the black carrot extracts [12].

#### *3.4. Antioxidant Activity*

The DPPH radical scavenging activity assay was performed according to several methods described previously [22,36–38] in which radical scavenging activities were determined by testing the extracts with the free radical DPPH and monitoring their absorbance decrease at 515 nm using a 1600-PC spectrophotometer (VWR, Graumanngasse, Vienna) with a 1 cm path length glass cells. Control assays using the black carrot extracts were performed in order to obtain their absorbance contributions. A solution of 50 µM DPPH was prepared using buffered methanol, which was prepared by mixing methanol with acetic acid buffer solution (0.1 M, pH 5.5) [36]. Then, 2.85 mL of DPPH solution were mixed with 150 µL of each extract and were left reacting 30 minat room temperature (~24 ◦C). Antiradical activity was expressed as Trolox equivalents per gram of fresh weight (µM TE g·fw−<sup>1</sup> ), which was calculated from the equation obtained using a linear regression after plotting the known absorbance with different Trolox concentrations, from 1 to 800 µM and *r* <sup>2</sup> = 0.9657.

A cupric reducing antioxidant capacity (CUPRAC) solution [24,26] was prepared to determine antioxidant capacity via electrochemical tests using CuCl<sup>2</sup> with a concentration of 3 mM in distilled water. Also, a solution of neocuproin at 6 mM in ethanol was prepared. In order to control pH of the main solution, a 1.2 M ammonium acetate buffer solution was prepared (pH = 7), then pH was adjusted adding 1.2 M HCl and 1.2 M NaOH as required. Concentrations of Trolox were varied from 1 to 800 µM in ethanol for obtaining a Trolox standard curve. 2 mL of each CuCl2, neocuproin, ammonium acetate buffer, Trolox and distilled water solutions were prepared and mixed. The 10 mL solution was stirred for 15 minand N<sup>2</sup> was bubbled into it for 5 min. The same procedure was followed to evaluate antioxidant capacity adding anthocyanins extracts instead of Trolox solution and the values are reported as a comparison between the analytical response vs. concentration plot during the antioxidant quantification because the slope is dependent on the stochiometric relationship between the antioxidant-oxidant species involved, which is related to the electron transfer per molecule pair.

The electrochemical tests were performed in a three-electrode electrochemical cell. A calomel Hg/Hg2Cl<sup>2</sup> (saturated with KCl) was used as reference electrode, a graphite bar was used as counter electrode and a glassy carbon (3 mm) electrode was used as working electrode. Before each electrochemical measurement, the working electrode was polished with aluminum oxide powder followed by ultrasonic stirring during 10 min; this process was repeated 3 times. The voltammograms were obtained in a Bio-LogicVP-50 potentiostat (Bio-Logic Science Instruments, Seyssinet-Pariset, France) with a sweep velocity of 100 mV s−<sup>1</sup> , starting the voltammograms from the open circuit potential (OCP) that was determined when the potential did not show a variation higher than 1 mV per second.

#### *3.5. Extract and Food Models Color Determination*

A custom MATLAB script (Mathworks Inc., Natick, MA, USA) was used to measure lightness and chromaticity coordinates in the L\* a\* b\* color space (CIELAB) according to CIE standard illuminant A (typical, domestic, tungsten-filament lighting with correlated color temperature of 2856 K). L\* indicates lightness, a\* and b\* are chromaticity coordinates, h (hue), c (chroma) and ∆E (color change) were calculated from a\* and b\* values. Digital images from samples were taken using a Sony digital camera α99II coupled with a Vario Sonnar T\* 24–70 mm lens (Sony Corporation, Tokyo, Japan) under the same light conditions; the images were cropped to 1024 × 1024 pixels and then processed with the afore mentioned script. Additionally, samples were measured in two different colored food models: yogurt and jelly. For yogurt food model, 10 g of commercial yogurt (Yoplait natural yogurt, Sigma Alimentos Lácteos México, Queretaro, Mexico) were colored using 10 mg of calculated anthocyanins from each extract and jelly was prepared using jelly powder (Coloidales Duche, Ciudad de México, Mexico) dissolved in boiling water (1:3 ratio) and 10 g of the mixture were colored using the same calculated amount of anthocyanins from each extract. Samples were measured at the beginning of the experiment and after 25 days, and compared with a colored yogurt/jelly with 10 mg of Red FD&C (Red Currant 12.5%, Colores Duche, Ciudad de México, Mexico) under the same conditions.

#### **4. Conclusions**

Anthocyanins were extracted from black carrot with ethanol/citric acid and ethanol/acetic acid for comparison between total anthocyanin content, and stability against media changes and antioxidant capacity was obtained to analyze samples in food models. Microencapsulation with TEOS was performed with the objective of enhancing anthocyanin stability. Extracts had the highest degradation in alkaline pH, and BCAR was the most stable sample to pH media changes. The antiradical activity of black carrots extracts was higher than other reported values, and when anthocyanin content is higher, the antioxidant effect increases. Results of UV radiation and thermal stability tests indicate that TEOS microencapsulation provides a negligible improvement in anthocyanins' stability. In conclusion, extraction with ethanol/acetic acid is the most convenient and stable treatment against pH media changes. Purification with resin and TEOS microencapsulation did not increase stability of the black carrot extracts. TEOS microencapsulation has proven to be effective (enhancing stability) for some dried materials from natural extracts in our previous research, but we do not recommend its use for materials obtained from black carrot extracts. Even though anthocyanins are already used in the food industry in beverages, our samples were not suitable for the yogurt or jelly model selected except for BCA sample in jelly that has the highest antioxidant activity, this gives it potential for being a functional natural colorant in this specific food model.

**Author Contributions:** Data and experimentation, G.A.M. and A.R.H.-M.; Formal analysis, R.E., A.R.H.-M. and M.E.; Funding acquisition, M.E.; Investigation, G.E.-A., I.S.-G. and J.M.-C.; Methodology, J.M.-C. and A.R H.-M.; Supervision, A.R.H.-M. and M.E.; Validation, A.L.R.-J.; Writing—original draft, G.A.M., A.R.H.-M. and M.E.; Writing—review & editing, A.L.R.-J.

**Funding:** This research received no external funding.

**Acknowledgments:** Authors would like to acknowledge the grant obtained from the Dirección General de Asuntos del Personal Académico (DGAPA) of Universidad Nacional Autónoma de México (UNAM) through "Programa de Apoyo a Proyectos de Investigación e Innovación Tecnológica" (PAPIIT No. IN209517). One of the authors, MSc Guillermo Espinosa-Acosta—fellow number 388907-; acknowledges the financial support received from CONACYT at the first period of the 2015 National Scholarships Call. Also, the authors are especially grateful to Perkin Elmer Mexico for obtaining the chromatograms shown in Figure 2 of this manuscript. Finally, authors are grateful to Bernardino Rodriguez-Morales and Adrian Hendrik Oskam Voorduin for their technical support; and also, to Gerardo Fonseca, and José Antonio Pérez Guzman, for their valuable comments about the manuscript, and to Angel Luis Rodríguez for his support on the development of the MATLAB script used for color analysis.

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

#### **References**

1. Hernández-Martínez, A.R.; Torres, D.; Molina, G.A.; Esparza, R.; Quintanilla, F.; Martínez-Bustos, F.; Estevez, M. Stability comparison between microencapsulated red-glycosidic pigments and commercial fd&c red 40 dye for food coloring. *J. Mater. Sci.* **2017**, *52*, 5014–5026. [CrossRef]


**Sample Availability:** Samples of the compounds are available from the authors.

© 2018 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Betanin, a Natural Food Additive: Stability, Bioavailability, Antioxidant and Preservative Ability Assessments**

**Davi Vieira Teixeira da Silva, Diego dos Santos Baião, Fabrício de Oliveira Silva, Genilton Alves, Daniel Perrone, Eduardo Mere Del Aguila and Vania M. Flosi Paschoalin \***

Instituto de Química, Universidade Federal do Rio de Janeiro, Av. Athos da Silveira Ramos 149, 21941-909 Rio de Janeiro, Brazil; daviufrj@outlook.com (D.V.T.d.S.); diegobaiao20@hotmail.com (D.d.S.B.); silvafo@live.com (F.d.O.S.); geniltonalves@gmail.com (G.A.); perrone@iq.ufrj.br (D.P.); emda@iq.ufrj.br (E.M.D.A.)

**\*** Correspondence: paschv@iq.ufrj.br; Tel.: +55-21-3938-7362; Fax: +55-21-3938-7266

Academic Editors: Lillian Barros and Isabel C.F.R. Ferreira Received: 11 January 2019; Accepted: 27 January 2019; Published: 28 January 2019

**Abstract:** Betanin is the only betalain approved for use in food and pharmaceutical products as a natural red colorant. However, the antioxidant power and health-promoting properties of this pigment have been disregarded, perhaps due to the difficulty in obtaining a stable chemical compound, which impairs its absorption and metabolism evaluation. Herein, betanin was purified by semi-preparative HPLC-LC/MS and identified by LC-ESI(+)-MS/MS as the pseudomolecular ion *m*/*z* 551.16. Betanin showed significant stability up to −30 ◦C and mild stability at chilling temperature. The stability and antioxidant ability of this compound were assessed during a human digestion simulation and ex vivo colon fermentation. Half of the betanin amount was recovered in the small intestine digestive fluid and no traces were found after colon fermentation. Betanin high antioxidant ability was retained even after simulated small intestine digestion. Betanin, besides displaying an inherent colorant capacity, was equally effective as a natural antioxidant displaying peroxy-radical scavenger ability in pork meat. Betanin should be considered a multi-functional molecule able to confer an attractive color to frozen or refrigerated foods, but with the capacity to avoid lipid oxidation, thereby preserving food quality. Long-term supplementation by beetroot, a rich source of betanin, should be stimulated to protect organisms against oxidative stress.

**Keywords:** beetroot; betalains; semi-preparative RP-HPLC; in vitro human gastrointestinal digestion; ex vivo colon fermentation; antioxidant ability; malonildialdehyde

### **1. Introduction**

Beetroot (*Beta vulgaris L*.) is a vegetable presenting significant scientific interest, mainly because it is a rich source of nitrate (NO<sup>3</sup> <sup>−</sup>), a compound with beneficial cardiovascular health effects, through the endogen production of nitric oxide (NO) [1]. Moreover, beetroots are the main source of betalains, a heterocyclic compound and water-soluble nitrogen pigment, which can be subdivided into two classes according to their chemical structure: betacyanins, such as betanin, prebetanin, isobetanin and neobetanin, responsible for red-violet coloring, and betaxanthins, responsible for orange-yellow coloring, comprising vulgaxanthin I and II and indicaxanthin [2,3]. Betalains are present in the tuberous part of beetroots, conferring its red-purple coloration.

Betanin (betanidin 5-*O*-β-D-glucoside) is the most abundant betacyanin and the only one approved for use as a natural colorant in food products, cosmetics and pharmaceuticals, under code EEC No. E 162 by the European Union and under Section 73.40 in Title 21 of the Code of Federal Regulations (CFR) stipulated by the Food and Drug Administration (FDA) in the United States [4–6] (Supplementary File—Figure S1A).

In the food industry, synthetic antioxidants are added to foods containing fat, especially meats, with the purpose of delaying oxidative processes that result in undesirable sensorial changes, decreased shelf life and nutritional value and the formation of secondary compounds potentially harmful to health [7,8]. However, data in the literature have associated the synthetic antioxidants BHT (butylated hydroxytoluene) and BHA (butylated hydroxyanisole) with possible deleterious health effects, as they have been reported as potential tumor promoters, following the chronic administration of these compounds to animals [9,10]. This has motivated the replacement of synthetic antioxidants with natural antioxidants extracted from foodstuffs [11,12].

Betanin can be used as a powerful antioxidant in the food industry in extract or powder form, and is also applied as a natural pigment. Its antioxidant activity in biological lipid environments has been demonstrated in human macromolecules, such as low density lipoproteins, membranes and whole cells [13]. Furthermore, betanin has attracted attention due to its anti-inflammatory and hepatic protective functions in human cells. This compound is able to modulate redox-mediated signal transduction pathways involved in inflammation responses in cultured endothelia cells, and has also displayed antiproliferative effects on human tumor cell lines [14,15]. In both healthy and tumoral human hepatic cell lines, betanin can induce the translocation of the erythroid 2-related factor 2 (Nrf2) antioxidant response element (ARE) from the cytosol to the nuclear compartment, which controls the mRNA and protein levels of detoxifying/antioxidant enzymes, including GSTP, GSTM, GSTT, GSTA (glutathione *S*-transferases), NQO1 (NAD(P)H quinone dehydrogenase 1) and HO– (heme oxygenase-1), in these cells, exerting hepatoprotective and anticarcinogenic effects [16].

Several betanin purification techniques have been reported, involving distinct steps and methodologies in order to purify this compound from vegetal sources, including complex food matrices such as beetroot. Among the methods employed for betanin purification, high-performance liquid chromatography (HPLC) and other chromatographic methods using reverse phase columns seem to provide the best balance between speed and efficiency [17]. However, no other studies have evaluated the stability of this molecule during storage conditions and its antioxidant capability after purification and during storage with the aim of use as a food additive.

In this context, the aim of the present study was to optimize a methodology applied for betanin purification in large amounts, using fresh juice obtained from red beetroot (*Beta vulgaris* L. species). In addition, the chemical stability and bioactivity of the purified molecule were also assessed, through two different viewpoints: (i) as a food preservative and (ii) as a *in natura* or processed food matrix component after consumption and simulated gastrointestinal and ex vivo colon fermentation processes.

#### **2. Results and Discussion**

#### *2.1. Betanin Purification*

The chromatographic profile of betanin and isobetanin in fresh beetroot juice was compared to a commercial standard (Figure 1A,B). In the fresh juice chromatogram, betanin corresponds to the major peak (RT = 18.17 min), followed by its isomer, isobetanin (RT = 19.27 min). Betanin is found in large amounts relative to its isobetanin isomer in fresh beetroot juice, which is similar to the ratio between betanin and isobetanin previously reported by Gonçalves et al. [17]. The betanin concentrations in fresh juice and in purified samples were calculated in comparison to a standard curve, described by the equation y = 5077078x − 370531 (R<sup>2</sup> = 0.9993). Linearity was obtained at betanin concentration ranging from 0 to 500 mg·mL−<sup>1</sup> . After purification an HPLC-diode array detector (DAD) analysis indicated a single peak at the maximum absorption λ (535–540 nm), characteristic for betacyanins (Figure 1C). Purified betanin can be preserved at 4 ◦C for at least 20 days and for 275 days if maintained at −30 ◦C (Figure 1D). On the contrary of what has been described previously [17], betanin preparations turned brown during shelf life, due to the possible action of polyphenol oxidases (PPO enzymes) in the

present study, whereas no sign of degradation was observed in the purified betanin. Temperature control, pH and non-exposure to light throughout the purification process reduced the chances of sample decomposition. Such precautions may have had a positive influence on the stability of the molecule, as demonstrated later in the stability assessment.

Purification technique yield is one of the most economically important aspects when obtaining natural food additives. The betanin extraction and purification yield from fresh beetroot juice and purified betanin recovered after chromatographic separation and mobile phase evaporation was calculated from an initial 500 g beetroot mass. The betanin concentration in juice was of 1.19 g mL−<sup>1</sup> and the amount of purified betanin recovered after purification by chromatographic separation and mobile phase evaporation was of 48 mg·mL−<sup>1</sup> corresponding to 4% of the initial betanin concentration. − ∙ −

As noted, in Figure 1C the beginning of betanin isomerization to isobetanin occurred, culminating with co-elution and a small base widening. The analytical column used for the separation and the elution conditions were not able to clearly separate the peaks in the initial isomerization stage, but the presence of the already formed isobetanin was well-evidenced and the peaks were clearly separated during storage (Figure 1D). It is noteworthy that betanin isomerization can be considered a structural modification rather than a degradation reaction. Isobetanin (2*S*/15*R*) differs from betanin (2*S*/15*S*) by the spatial conformation of the carbonyl group at carbon 15, but exhibits similar chromatic properties to betanin with no observable color change [18].

**Figure 1.** Betanin separation by high-performance liquid chromatography diode array detector (HPLC-DAD) monitored at 536 nm. (**A**) Betanin standard chromatographed in the analytical HPLC column, (**B**) fresh beetroot juice sample chromatographed in semi-preparative HPLC, (**C**) betanin purified by semi-preparative HPLC and separated using an analytical HPLC column and (**D**) betanin evaluated after 275 days of freezing and chromatographed using an analytical HPLC column. Betanin (peak 1) and isobetanin (peak 1'). The betanin chemical structure from red beet was reproduced from Cai et al. [19].

#### *2.2. HPLC-ESI(+)-MS/MS Analysis*

HPLC-purified betanin identification was performed by mass spectrometry, as displayed in Figure 2A,B. The pseudomolecular ion *m/z* 551.16 corresponding to betanin (Figure 2A) and its characteristic fragmentation (Figure 2B), the ion *m*/*z* 389.11 corresponding to its aglycone form, betanidin, a precursor structure of betacyanins (Supplementary File—Figure S1B), were observed, corroborating previous findings reported by Gonçalves et al. [17] and Netzel et al. [20]. ′

**Figure 2.** Identification of purified betanin by HPLC-ESI(+)-MS/MS. (**A**) betanin *m*/*z* 551 [M + H]<sup>+</sup> , (**B**) fragmentation of purified betanin *m*/*z* from the MS/MS of 551 [M + H]<sup>+</sup> .

− ∙ − ∙ − It can be suggested that the *m*/*z* 637 adduct may be the result of the condensation of the formic acid (molecular weight 46) used in betanin purification and a decarboxylated betanin derivative, forming 6 ′ -*O*-malonyl-2-decarboxyl-betanin (molecular weight 592), a decarboxylated betanin derivative [21]. The mass spectrometry analysis confirmed that the purification procedure was successful in isolating betanin in its purified form.

#### *2.3. Storage Stability*

∙ − ∙ − ∙ − Betanin was stable for over 275 days (9 months) when stored at −30 ◦C at pH 7 (Figure 1D) and at least for 20 days when stored at 4 ◦C. No significant loss of the betanin samples was observed before and after frozen storage (21.30 ± 1.98 mg·mL−<sup>1</sup> versus 17.23 ± 4.82 mg·mL−<sup>1</sup> , *p* < 0.001, respectively). The initial concentration of purified betanin samples during storage at refrigerator temperature was unaltered during the first 20 days (12.13 ± 0.70 mg·mL−<sup>1</sup> versus 13.51 ± 1.01 mg·mL−<sup>1</sup> , *p* < 0.001) but on the 27th day, a loss of 25% of the initial betanin concentration was observed (9.72 ± 0.60 mg·mL−<sup>1</sup> ).

ff − Most betalains, including betanin, are under-utilized as colorants in processed foods due to reports concerning poor stability compared with the shelf-life of foods. In standard storage conditions betacyanin stability in spray dried beetroot powder was reported by Kaimainen et al. [22], who assessed the product stability by HPLC at 535 nm and pH 5 during storage for 25 weeks at different temperatures, including frozen (−20 ◦C) and chilled (4 ◦C). Betacyanins in beetroot powder remained unchanged during 4 months under freezing. However, stability under refrigeration was not well established. It is noteworthy that in that study, beside the addition of sweeteners, betanin was not in its purified form, but rather protected by the food matrix that naturally contains antioxidants and chelating agents, which may exert protective effects on the chemical structure of betanin maintained under freezing conditions [18]. Factors such as temperature, pH, type of buffer solution and the presence or absence of oxygen can affect betanin stability during storage. Betanine degradation results in the formation of betalamic acid and cyclo-dopa-5-*O*-glycoside. However, betanin can also display the ability to degrade and regenerate continuously during storage, as the reaction is reversible, thus maintaining betanin concentrations [23]. This continuous betanin regeneration capacity during the storage process is still not well-elucidated in the literature [2,23,24]. Therefore, the stability results found in the present study indicate that betanin, if used as an additive in refrigerated or frozen foods, would remain stable

during the time and temperature recommended by the legislation as ideal for preserving meats and meat-derivatives from 6 to 12 months at temperatures below −18 ◦C and for 5 days at 4 ◦C [25]. −

#### *2.4. Lipid Peroxidation Inhibition in Meat Matrices*

Betanin's ability to inhibit lipid peroxidation process in meat was assessed by thiobarbituric acid reactive substance (TBARS) determination. Control samples (without antioxidants) showed the highest malondialdehyde (MDA) concentrations when compared to samples treated with betanin, BHA or BHT during 9 days of storage, determined on the 3rd, 6th and 9th days. Betanin 2% was equally effective in inhibiting lipid peroxidation when compared to the synthetic antioxidants BHA and BHT on the 3th and 6th day of storage (*p* < 0.05). Although the amounts of MDA in meat samples treated with betanin 2% (*w*/*w*) (5.07 ± 0.03 mg·kg−<sup>1</sup> ) were 22% and 16% higher than those found in samples treated with BHA (4.47 ± 0.10 mg·kg−<sup>1</sup> ) and BHT (4.28 ± 0.38 mg·kg−<sup>1</sup> ) on the 9th day of storage, the meat samples treated with betanin 2% (*w*/*w*) still presented lower MDA concentrations than the samples with no preservative addition (7.15 ± 0.07 mg·kg−<sup>1</sup> vs 5.07 ± 0.03 mg·kg−<sup>1</sup> ) (Figure 3). ∙ − ∙ − ∙ − ∙ − ∙ −

**Figure 3.** Lipid oxidation in ground pork loin evaluated by the production of malondialdehyde (MDA) during 9 days of storage at 4 ◦C. Control H2O-DD, BHA (buthylated hydroxyanisole), BHT (butylated hydroxytoluene), Betanin 2% (*w*/*w*). Data are expressed as the means ± SD of three independent determinations. Different letters indicate differences between days at a significance level of *p* < 0.01. The symbol \* (*p* < 0.05) indicates differences compared to day 0. The symbol \*\* (*p* < 0.05) indicates differences compared to day 3.

Few studies assessing the effect of the addition of beetroot or beetroot-extracted compounds on the oxidative stability of foods susceptible to lipid oxidation are available. The effect of beetroot inclusion as a mayonnaise ingredient promoted a higher inhibitory effect on lipid oxidation compared to the commercial antioxidant [26]. Contradictory results, however, were observed in fermented meat sausages, since no beetroot effect was observed [27].

To the best of our knowledge, the present study is the first to evaluate the use of purified betanin as a natural antioxidant in food matrices. Lipid oxidation is one of the main factors affecting food quality and is directly related to nutritional value and sensorial characteristics. The present study indicates that betanin used as an additive at the concentration of 2% (*w*/*w*) is a potential substitute for synthetic antioxidants in the preservation of refrigerated meat. Furthermore, betanin can exert its maximum protective effect against lipid oxidation for 6 days exceeding the 5-day shelf-life recommended for refrigerated meats [25].

#### *2.5. Betanin Chemical Stability during In Vitro Simulated Gastrointestinal Digestion*

A 23 mg·mL−<sup>1</sup> dose was used to assess betanin bioavailability during in vitro human simulated gastrointestinal digestion by continuous multistage steps. A small loss was observed in betanin content after the oral phase digestion. However, more important decreases were observed after the gastric simulated digestion, reaching 65% of the initial sample content, and lowering to 46%, after small intestine simulated digestion (Table 1). No betanin was detected after the ex vivo colon fermentation assay, where the remaining betanin recovered at the end of the in vitro simulated gastrointestinal digestion, corresponding to 54% of the original sample, was assayed by ex vivo colon fermentation (Supplementary File—Figure S2).

**Table 1.** Betanin concentrations during in vitro simulated gastrointestinal digestion.


Betanin availability was determined by reverse phase high-performance liquid chromatography diode array detector (RP-HPLC-DAD), assessed through changes in the peak area determined at 536 nm. In vitro human gastrointestinal digestion was sequentially simulated and samples were harvested at each phase. The ex vivo colon assay was performed incubating the digested material obtained after the entire in vitro gastrointestinal digestion with fresh feces donated by seven healthy volunteers. Data are expressed as the means ± SD of three independent experiments. Different letters in the same line indicate significant differences between samples (*p* < 0.01).

Only a few studies presenting limitations are available on the chemical stability of the purified betanin in in vitro simulated digestion through the gastrointestinal tract. In a previous report, betanin was degraded by 75% and 35% after the gastric and intestinal phases, respectively [28]. However, the sample was added directly to each fluid—gastric or intestinal, generating no information about physiological digestion in the digestive tract. In the present study, an in vitro simulated digestion was conducted sequentially, where betanin was added to simulated oral fluid and digestion was observed by a sequential in vitro system, transferring aliquots to the simulated gastric fluid, resulting in a decrease of 35% after gastric digestion. This 35% decrease in betanin contents observed after gastric digestion is due to its impaired stability at acidic pH 2. It is known that betalains exhibit stability at pH ranging from 3 and 7 [18]. A significant decrease in betacyanin stability in a solution containing hydrochloric acid at pH 2.0 at 37 ◦C was observed, whereas betacyanins maintained at pH 4.7 were less susceptible to degradation [29]. In acid pH, the betanin structure can be degraded in C-17 decarboxylation, dehydrogenation and cleavage of betalamic acid and cyclo-Dopa-5-*O*-βglycoside [2,30]. Herein, in addition to exposure to acidic pH, the absence of the food matrix may have exacerbated the betanin susceptibility to gastric fluid degradation, since it has been previously demonstrated that betanin and its isobetanin isomer (unpurified) can be protected from stomach digestion by the food matrix [31].

In addition, an overall decrease in betanin content to approximately 46% was noted when the simulated gastric fluid digestion was performed, followed by an 11% decrease during intestinal digestion. Besides the absence of the protective effect of the food matrix, the influence of pH on betanin stability is reinforced by the data reported herein, comparing the percentage of loss in the gastric phase to the intestinal phase (35% vs 11%). Small intestine pH is around 6.5, matching the reported pH-range of betanin stability and corroborating the lower betanin degradation at the small intestine [31].

Several polyphenols are described as reaching the large intestine, where they are absorbed following metabolization by colon bacteria consortia [32]. In the present study, betanin was detectable by HPLC at the 536–540 nm range, while no other metabolite derived from its biosynthetic conversion was observed. Although betanin was shown to be stable at 30 ◦C and 4 ◦C, the 24 h exposure to 37 ◦C and environmental conditions in the colon lumen may still promote its degradation. In a murine model study, only 3% of betanin administered to animals were recovered in feces after 24 h, indicating that colon absorption is not likely [28]. Permeability and solubility are important barriers concerning colon absorption in humans that, alongside the metabolic activity of bacterial consortia and the mild temperatures within the large intestine lumen point to improbable colon absorption and/or maintenance of the chemical stability of the betanin molecule.

#### *2.6. Betanin Antioxidant Activity throughout Simulated Human Gastrointestinal Digestion*

Betanin in its purified form was able to inhibit the OH-radical in the total antioxidant potential (TAP) assay. The OH-radical is considered the most reactive oxidant in living organisms, generated by the Fenton reaction [33] (Supplementary File—Figure S3). In the ferric reducing ability of plasma (FRAP) assay, betanin was effective in reducing the ferric ion of the tripyridyltriazine complex (Fe3+-TPTZ) to the ferrous ion (Fe2+-TPTZ), reflecting its ability to donate electrons and reduce reactive species. In addition, betanin was effective in reducing the ABTS<sup>+</sup> radical, as observed in the trolox equivalent antioxidant capacity (TEAC) and oxygen radical antioxidant capacity (ORAC) assays (Table 2). In addition, betanin showed a high TAP value after both oral and intestinal digestion.


**Table 2.** Total betanin antioxidant potential and antioxidant activity pre and post in vitro simulated human gastrointestinal digestion.

Betanin antioxidant potential and antioxidant activity were evaluated before and after the simulated human gastrointestinal digestion using different assays, namely FRAP, TEAC and ORAC. Data are expressed as the means ± SD from three independent experiments. Different letters in the same column indicate difference at a significance level of *p* < 0.001.

The high antioxidant activity of betanin is well-documented [34]. However, to promote health beneficial in human beings, the chemical structure of the ingested betanin and its antioxidant properties should be maintained in the gastrointestinal absorption site. The antioxidant activities of purified betanin following the final sequential digestive process through the gastrointestinal apparatus increased when compared to pre-digested samples, as demonstrated in the FRAP and TAP assays (Table 2).

When assessing each digestion fluid, the antioxidant activity of betanin evaluated by the FRAP assay was increased in the post-oral digestion assays. The FRAP assay increments can be attributed to natural antioxidants originally present in human saliva [35]. After small intestine digestion, purified betanin increased or maintained the antioxidant activities evaluated by the TAP, FRAP, TEAC and ORAC assays when compared to the oral digestion processes (Table 2).

Additionally, the pH of the different fluids in the human body undergoes variations, influencing the stability and bioactivity of betanin in the different digestive tract compartments. Salivary fluid present a pH of about 7.4, whereas stomach is maintained between 1.5 and 3.5, while the abdominal cavities, including the small and large intestine, display a pH of 7.4.

The antioxidant ability of betanin was reduced in the acidic pH of the gastric fluid, evidencing the pH-dependence of the free radical-scavenging activity of betanin, but, countering its antioxidant activity in the alkaline environment of the small intestine lumen, where the antioxidant activity assessed by all assays was either increased or maintained at the same level of the oral fluid, as mentioned previously (Table 2). The alternate antioxidant activity of betanin between low and high levels in the simulated gastric compartment and in the small intestine indicates that the chemical structure of betanin was maintained unaltered following acidic pH exposure during the simulated gastric digestion, but should be attributed to the protonation of the betanin molecule, favoring the maintenance of free radical-scavenging activity until absorption in the gastrointestinal tract.

Changes in pH fluids induce changes in the chemical structure of betanin along the gastroenteric apparatus. Betanin presents its cationic form at pH < 2, zwitterionic form at pH = 2, anionic mono at 2 < pH < 3 with deprotonated C2-COOH and C15-COOH groups, anionic bis at 3.5 < pH < 7 with C2-COOH, C15-COOH and C17-COOH groups deprotonated and anionic tris at 7.5 < pH < 9 with all carboxyl deprotonated groups, in addition to the C6-OH group on the phenolic ring (Supplementary File—Figure S4) [36]. The increase in betanin bioactivity, when in an alkaline pH environment, as found in the simulated small intestine fluid, can be ascribed to its ability to donate H<sup>+</sup> and electrons when altering from the cationic to the mono, bis and tri deprotonated states. The free radical scavenging activity of betanin at different pH (from 2 to 9) was previously determined through the TEAC assay, phenolic O–H homolytic bond dissociation energy (OH BDE), ionization potential (IP) and deprotonation energy (DE) [37]. The TEAC assay indicated that the antioxidant activity of betanin is dependent on pH, and very high above pH > 4. Moreover, with the gradual increase of the deprotonation of the betanin molecule (mono, bi- and tri-deprotonated) according to increasing pH, BDE and PI values decreased. This implies that, at slightly alkaline pH, betanin becomes a better hydrogen and electron donor, increasing its radical-scavenger ability as observed in the antioxidant assays in the simulated small intestine fluid when compared to the gastric fluid.

Although almost 46% (11 mg) of betanin content were chemically modified in the gastric tract, the antioxidant power of the remaining betanin, 54% of the original amount, corresponding to 12 mg found in the small intestine fluid (21 µmol) seems to be enough to promote lipid oxidation inhibition, since betanin in the range of 0.3–1.9 µmol has been found to inhibit lipid peroxidation in biological membranes, in a linoleate emulsion catalyzed by the "free iron" redox cycle, in H2O2-activated metmyoglobin and in lipoxygenase activity [38].

#### **3. Material and Methods**

#### *3.1. Standards and Reagents*

The betanin standard (C24H26N2O13), sulfuric acid (H2SO4), boric acid (H3BO3), formic acid (CH2O2), hydrochloric acid (HCL), terephthalic acid (TPA, C8H6O4), ethylenediaminetetraacetic acid (EDTA), 6-hydroxy-2-5-7-8-tetramethylchromo-2-carboxylic acid (Trolox), ascorbic acid (C6H8O6), sodium hydroxide (NaOH), potassium permanganate (KMnO4), hydrogen peroxide (H2O2), potassium sulphate (K2SO4), ferrous sulphate (FeSO4), methyl red, bromocresol green, petroleum ether, anhydrous sodium acetate (CH3COONa), tetrabutylammonium perchlorate (C16H36N.H2PO4), vanillin, tripyridyltriazine (TPTZ, C18H12N6), iron chloride (FeCl3), dibasic sodium phosphate (NaH2PO4.H2O), monobasic sodium phosphate (NaH2PO4.H2O), sodium chloride (NaCl), anhydrous monobasic sodium phosphate (Na2HPO4), sodium bicarbonate (NaHCO3) C-211,2,2′ -Azobis (2-methylpropionamidine), dihydrochloride (AAPH), 2,2′ -azinobis [3-ethylbenzothiazoline-6-sulfonic acid]-diammonium salt (ABTS, C18H24N6O6S4), sodium fluorescein (C20H12O5), potassium hydroxide (KOH), ammonium thiocyanate (NH4SCN), trichloroacetic acid (Cl3CCOOH) were purchased from Sigma-Aldrich Chemical Co. (São Paulo, SP, Brazil). Methanol (MeOH), ethanol, acetone, and acetonitrile were purchased from Tedia Company Inc. (Rio de Janeiro, RJ, Brazil). Buthylated hydroxyanisole (BHA), butylated hydroxytoluene (BHT), 1,1,3,3-tetramethoxypropane ((CH3O)2CHCH2CH(OCH3)2) and 2-thiobarbituric acid (C4H4N2O2S) were purchased from Sigma-Aldrich Co. HPLC grade Milli-Q water (Merck Millipore, Burlington, MA, USA) was used throughout the experiments.

#### *3.2. Betanin Purification*

#### 3.2.1. Sample Preparation

Red beetroot was peeled, sliced and homogenized using a centrifuge food processor EC 700 (Black and Decker, São Paulo, Brazil). The homogenates were centrifuged at 15,000× *g* for 30 min at 25 ◦C and filtered through a PTFE filter membrane 25 mm, pore size 0.45 µm (Merck-Millipore). The supernatants (4 mL) were concentrated under reduced pressure (18 mbar, 25 ◦C) and resuspended in 2 mL deionized water.

#### 3.2.2. HPLC Betanin Purification

Concentrated beetroot juice was purified by RP-HPLC. The HPLC apparatus consisted in an LC-20A Prominence, (Shimadzu®, Kyoto, Japan) equipped with a quaternary pump and a DAD model SPD-M20A (Shimadzu®, Kyoto, Japan). A 15 µm Phenomenex C18 column (250 × 21.2 mm I.D., Torrance, California, USA) connected to an FRC-10A fraction collector (Shimadzu®) was used in the semi-preparative HPLC. The elution conditions were performed according to Cai et al. [39] with modifications. Solvent A was 1% formic acid, and solvent B was 80% methanol at a linear gradient (0–25 min, 11–55%). The injection volume was 100 µL and a flow rate of 5.5 mLmin−<sup>1</sup> was used. Separations were monitored at 536 nm and, after purification, magenta fractions, containing betanin, were concentrated by a rotary evaporator (Rotavapor® R-215, Buchi, São Paulo, Brazil) at 24 ◦C, 150 rpm and a water bath at 40 ◦C. The extracts were then suspended in 1 mL deionized water and stored at −30 ◦C under an N<sup>2</sup> atmosphere for further analysis. The purified betanin was analyzed using a Nucleosil 100-C18 column (250 × 4.6 mm I.D., 5 µm) with 30 µL injection volume and a flow rate of 1.0 mL min−<sup>1</sup> . The mobile phase and gradient conditions were similar to the purification step and betanin concentrations were quantified in comparison to a betanin standard solution (Sigma-Aldrich Co.).

#### *3.3. Betanin Identification by Liquid Chromatography Positive Ion Electrospray Ionization Tandem Mass Spectrometry (LC-ESI(+)-MS/MS)*

Mass spectrometry was performed as described by Gonçalves et al. [17]. The RP-HPLC purified fraction was ionized in the positive mode and ions were monitored in the full scan mode (range of *m*/*z* 50–1500). The ESI(+)-MS/MS analysis was carried out on a Bruker Esquire 3000 Plus Ion Trap Mass Spectrometer (Bruker Co., Billerica, MA, USA) equipped with an electrospray source in the positive ion mode. Nitrogen was used as the nebulizing (45 psi) and drying gas (6 L·min−<sup>1</sup> , 300 ◦C) and helium as the buffer gas (4 × 10−<sup>6</sup> mbar). The high capillary voltage was set to 3500 V. To avoid space–charge effects, smart ion charge control (ICC) was set to an arbitrary value of 50.000. Betanin identification was based on its mass (550 g·mol−<sup>1</sup> ) and by similarity with the commercial standard and literature-available spectra [39].

### *3.4. Storage Stability*

The stability of purified betanin during refrigeration (4 ◦C) and freezing (−30 ◦C) was evaluated by RP-HPLC-DAD (Shimadzu®, Kyoto, Japan), monitoring changes in the area under the chromatogram peak obtained at 536 nm, in similar conditions as those described for the betanin analysis.

#### *3.5. Betanin Ability to Inhibit Lipid Peroxidation in Meat*

Betanin ability to inhibit lipid peroxidation was evaluated by MDA determination in meat TBARS assay, as described previously [40] with modifications. A sample of ground pork loin (500 g) from a local butcher shop in Rio de Janeiro, Brazil and divided into 4 portions and treated as follow: (i) ground pork loin non-treated by antioxidants; (ii) ground pork loin treated with betanin (2%; *w*/*w*); (iii) ground pork loin treated with BHT (0.01%); (iv) ground pork loin treated with BHA (0.01%). MDA extraction was performed in 3.0 g of each meat sample homogenized with 9 mL of 7.5% TCA. The homogenate was centrifuged at 3000× *g* for 15 min at 25 ◦C and filtered through Whatman n◦ 4 paper (Merck Millipore Co). TMP (the MDA standard) at 3.2 mM in 0.1 M HCl (stock solution) was kept for 2 h at room temperature in the dark. After hydrolysis, the TMP solution was diluted with 7.5% TCA to the concentrations of 1, 2, 4, 8, 16 and 32 µM. After, 1 mL of MDA at different concentrations or 7.5% TCA solution (blank) was transferred into a screw-cap tube and 1 mL of 20 mM TBA solution was added. The tubes were heated in a boiling water bath at 90 ◦C for 30 min and cooled in tap water for 10 min. Absorbance of the MDA-TBA adducts were measured at 532 nm on a spectrophotometer DU®530 (Beckman Coulter Inc., Brea, CA, USA). Because betanin absorbs light in the range of 530–540 nm, additional blanks containing betanin (1 or 2%), TCA or TBA (no meat) were used to correct the overestimation of the TBA-MDA adduct absorbance. The concentration of MDA was expressed in mg of MDA per kg of meat (mg of MDA·kg−<sup>1</sup> meat) at each treatment along the 9 days storage at 4 ◦C.

#### *3.6. TAP Determination*

Betanin samples were analyzed as described previously [41]. Samples were diluted (1:10) and centrifuged at 4500× g for 10 min, and the supernatants were then filtered through 0.45 µm cellulose membranes (Merck Millipore Co). The resulting samples were transferred to amber vials and incubated at 37 ◦C for 10 min with a solution containing 1 mM Fe2+, 10 mM H2O<sup>2</sup> and 1 mM terephthalic acid (TPA) in 50 mM phosphate buffer pH 7.4. Hydroxyterephthalic acids (HTPA) were detected by HPLC. TAP measurements were obtained by the difference between the chromatogram surface area generated in the Fenton reaction with and without the sample.

#### *3.7. Antioxidant Activity Determination by Different Assays*

#### 3.7.1. FRAP Determination

FRAP assays were performed using a modification of the method described by Benzie and Strain [42]. Betanin samples were diluted (1:10) and then mixed thoroughly with the FRAP reagent. Standard FeSO<sup>4</sup> solutions were used and absorbances at 593 nm were determined on a V–530 UV/VIS spectrophotometer (Jasco®, Easton, PA, USA). The FRAP results for each sample were evaluated in triplicate and expressed as µmol of Fe2+ ·L −1 .

#### 3.7.2. TEAC Determination

TEAC assays were performed using a modification of the method described by Re et al. [43]. The ABTS radical cation (ABTS•<sup>+</sup> ) was generated by chemical reaction of ABTS with K2S2O<sup>8</sup> in the dark at room temperature for 12–16 h. Each betanin sample was mixed with the ABTS•<sup>+</sup> reagent and absorbances at 720 nm were determined using a V–530 UV/VIS spectrophotometer (Jasco®). TEAC results were determined in triplicate and were associated to the ABTS•<sup>+</sup> inhibition percentage by antioxidants present in the samples. The TEAC results for each beetroot sample were evaluated in triplicate and expressed as µmol of Trolox·L −1 .

#### 3.7.3. ORAC Determination

The ORAC assay was performed according to Zuleta et al. [44], with modifications. Sample absorbances were determined on a Wallac 1420 VICTOR multilabel counter (Perkin–Elmer Inc, Waltham, MA, USA) with fluorescence filters at an excitation wavelength of 485 nm and emission wavelength of 535 nm. A fluorescein stock solution was prepared by weighing 3 mg of fluorescein followed by dissolution in 100 mL of phosphate buffer (75 mM, pH 7.4). The fluorescein stock solution was stored in complete darkness under refrigeration. The fluorescein working solution (78 nM) was prepared daily by dilution of 0.100 mL of the fluorescein stock solution in 100 mL of phosphate buffer. The AAPH radical (221 mM) was prepared daily by mixing 600 mg of AAPH in 10 mL phosphate buffer. A 25 µM Trolox solution was used as reference standard, prepared daily in phosphate buffer from a 4 mM stock standard solution kept in a freezer at 20 ◦C. A total of 100 µL of fluorescein (78 nM)

and 100 µL of the samples, blanks (phosphate buffer), or standards (25 µM of Trolox) were added to each well, followed by 50 µL of AAPH (221 mM). ORAC values, expressed as µM Trolox equivalents were calculated by applying the following formula:

ORAC (µM Trolox equivalents) = CTrolox·(AUCSample − AUCBlank)·*k*

$$\text{(AUC}\_{\text{Troloc}} - \text{AUC}\_{\text{Blank}})$$

where CTrolox is the Trolox concentration (µM), *k* is the sample dilution factor, and AUC is the area below the fluorescence decay curve of the samples, blanks and Trolox, respectively, calculated using the GraphPad Prism v.5 software package (GraphPad Software Inc., San Diego, CA, USA). ORAC determinations were performed in triplicate and expressed as mmol Trolox equivalents·100 g−<sup>1</sup> .

#### *3.8. Simulated Betanin In Vitro Human Gastrointestinal Digestion and Ex Vivo Colon Fermentation (Supplementary File—Figure S5)*

Betanin concentrations after in vitro oral, gastric and small intestine digestion and ex vivo colon fermentation were evaluated by RP-HPLC while antioxidant activity was evaluated by TAP, FRAP, TEAC and ORAC assays, as described previously. Samples were analyzed in triplicate.

The in vitro human simulated gastrointestinal digestion, including the oral, gastric and small intestine phases, was performed according to Oomen et al., [45] and Sagratini et al. [46], with modifications. For the OD simulation, 1 mL betanin aliquots at 23 mg·mL−<sup>1</sup> were placed in a glass jar followed by the addition of 3 mL of human saliva, and incubated at 37 ◦C for 1 min under orbital agitation at 260 rpm in a Sorvall ST 16R centrifuge (Thermo ScientificTM, Waltham, MA, USA) to complete the OD.

A 2.5 mL aliquot of artificial gastric fluid containing 2.75 g of NaCl, 0.27 g of NaH2PO4, 0.82 g of KCl, 0.42 g of CaCl2, 0.31 g of NH4Cl, 0.65 g of glucose, 0.085 g of urea, 3.0 g of mucine, 2.64 g of swine gastric pepsin, 1.0 g of bovine albumin, 8.3 mL of HCl was added to the oral fluid sample to a final volume of 500 mL and the pH was adjusted to 2.0 with 5 M HCl. The glass jars were then resealed with a rubber septum and the atmosphere was saturated with N<sup>2</sup> and incubated at 37 ◦C for 2 h under orbital shaking at 260 rpm to complete the GD.

The gastric fluid had its pH adjusted to 6.0 with NaHCO<sup>3</sup> and 2.0 mL of artificial small intestine fluid containing 6.75 g of NaCl, 0.517 g of KCl, 0.205 g of CaCl2, 3.99 g of NaHCO3, 0.06 g of KH2PO4, 0.0375 g of MgCl2, 0.1375 g of urea, 25.0 g of swine bile, 4.0 g of swine pancreatin, 1.2 g of albumin bovine and 0.185 mL HCl were added to a final volume of 500 mL. The glass jars were then resealed and the atmosphere was saturated with N<sup>2</sup> and incubated at 37 ◦C for 2 h under orbital shaking at 260 rpm to complete the ID. At the end of each simulated gastrointestinal step (oral, gastric and small intestine), aliquots were collected and centrifuged (3000× *g*, 15 min, 25 ◦C). The supernatants were then filtered through 0.45 and 0.22 µm membranes, followed by Amicon ultra filtration using 10 kDa cut-off membranes.

#### Ex vivo Colon Fermentation

The ex vivo colon fermentation assay was performed according to Hu et al. [47] with modifications, in accordance to the ethical standards of the declaration of Helsinki after approval by the Hospital Universitário Clementino Fraga Filho/Universidade Federal do Rio de Janeiro Education and Research Committee, under No. 512.84.

The ex vivo assay was performed using fresh feces donated by seven healthy volunteers (4 men and 3 women), recruited according to the following criteria: age between 18 and 50, eutrophic (BMI between 18.5 and 24.9 kg m<sup>2</sup> ), absence of gastrointestinal diseases, displaying one bowel movement every two days and up two bowel movements per day, with no medication and/or food supplements used 90 days prior to the feces collection.

The feces were homogenized in a nutrient-rich medium (0.5 g·10 mL−<sup>1</sup> ), prepared according to McDonald et al. [48], where the medium was autoclaved and saturated with CO<sup>2</sup> in an anaerobic chamber for 48 h. A 5 mL aliquot of this mixture was added to the digested material after the in vitro gastrointestinal digestion. The mixture was then incubated at 37 ◦C, under orbital shaking at 50 rpm for 48 h. The ex vivo colon fermentation assay was independently repeated three times.

#### *3.9. Statistical Analyses*

A one-way analysis of variance (ANOVA) with repeated measurements were performed to identify differences in TAP and antioxidant activities (FRAP, TEAC and ORAC) before and after the simulated gastrointestinal digestion (pre-digestion, oral, gastric, small intestine and colon phases). In addition, a two-way analysis of variance (ANOVA) with repeated measurements was performed to identify differences in MDA concentrations in the lipid peroxidation assay between each type of antioxidant and between each experiment day, evaluated by TBARS assay. When a significant *F* was found, an additional *post hoc* analysis was performed by a Bonferroni correction. Data were expressed as means ± standard deviation (SD). The statistical analyses were performed using Graphpad Prism software version 5 for Windows® (GraphPad Software, San Diego, CA, USA).

#### **4. Conclusions**

The decomposition of food components and bioactive or additive compounds along the gastrointestinal trait is a helpful tool to assess their potential positive and negative effects on health, as well as to evaluate possible toxicity and/or safety usage. The prediction of the in vivo absorption of such compounds can aid in compound usage regulation, establishing safety dosages in food.

Betanin in its purified form can be very stable during storage at low temperature and alkaline pH, so it may be useful as food colorant and antioxidant additive in meat and meat derivatives as a substitute for synthetic antioxidants. According to the chemical stability at both refrigerator and freezer temperatures, betanin can be used as a food colorant or preservative in almost all foodstuffs, including those stored at −30 ◦C, such as bacon, sausages, vegetables, ham, corned beef, ice cream and sherbet (www.fda.gov/food/guidance/regulation) and those preserved at 4–8 ◦C, including minced meat and fresh meat, yogurts and desserts, since their shelf life is lower than the 20 days in which betanin is stable.

In addition, betanin maintained its bioactivity during the simulated digestive process, presenting high TAP, FRAP, TEAC and ORAC values in the intestinal phase. Although the exact parts of the gastrointestinal tract in where betanin is absorbed still require elucidation, it can be suggested that absorption may occur in the small intestine. The chemical integrity of betanin and its antioxidant activity can be considered potential aid against diseases caused by oxidative stress.

These novel findings reinforce the importance of the regular uptake of red beetroot and its derivative products. The formulation of new dietary supplements or processed foods can include purified betanin, not only as a natural food colorant or preservative, but also as a bioactive compound that may act as an adjuvant in the treatment and prevention of chronic diseases related to oxidative stress in humans.

**Supplementary Materials:** Figure S1: Chemical structure of betanin (A) and betanid in (B); Figure S2: Betanin chromatograms before and after each in vitro digestion phase assessed by RP-HPLC equipped with DAD detector (536 nm). Purified betanin (A), after oral digestion (B), after gastric digestion (C), after small intestine digestion (D); Figure S3: Betanin total antioxidant potential (TAP) assessed by RP-HPLC equipped with a fluorescence detector (312/428 nm). Hydroxyterephthalic acid (HTPA) chromatograms of generated in the Fenton reaction without any sample (A), after betanin addition (B), after oral digestion (C), after gastric digestion (D), after small intestine digestion (E); Figure S4: Influence of pH on betanin chemical structure charge changes in an aqueous solution according to Frank et al. [35]. Figure S5: Simulated digestion scheme.

**Author Contributions:** D.V.T.d.S., E.M.D.A. and V.M.F.P. conceptualized and designed the research; D.V.T.d.S., D.d.S.B., F.d.O.S., G.A. and D.P. compiled, quantified and performed all experiments; D.V.T.d.S., D.d.S.B., E.M.D.A. and V.M.F.P. evaluated and interpreted all the results; D.V.T.d.S. and D.d.S.B. prepared the figures; D.V.T.d.S and V.M.F.P. wrote the manuscript; D.V.T.S., D.d.S.B. and V.M.F.P. edited and revised the manuscript. All authors critically revised the manuscript concerning important intellectual content, and read and approved the final manuscript.

**Funding:** The authors acknowledge financial support from CNPq (Conselho Nacional de Desenvolvimento Científico e Tecnológico), CAPES (Coordenação de Aperfeiçoamento de Pessoal de Nível Superior) and FAPERJ (Fundação Carlos Chagas Filho de Amparo à Pesquisa do Estado do Rio de Janeiro).

**Conflicts of Interest:** This article does not report any studies with human or animal subjects. The authors declare no conflicts of interest.

#### **References**


**Sample Availability:** Samples of the compounds are available from the authors.

© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Influence of Oxygen-Containing Sulfur Flavor Molecules on the Stability of** β**-Carotene under UVA Irradiation**

#### **Gong-Liang Zhang 1 , Hong-Yan Wu 2 , Ying Liang 2 , Jie Song 1 , Wei-Qi Gan <sup>1</sup> and Hong-Man Hou 1, \***


#### Academic Editor: Lillian Barros

Received: 28 December 2018; Accepted: 13 January 2019; Published: 16 January 2019

**Abstract:** The influence of 11 kinds of oxygen-containing sulfur flavor molecules was examined on β-carotene stability under UVA irradiation in ethanol system. Both the effects of sulfides on dynamic degradation of β-carotene and the relation between structure and effect were investigated. The oxidation products of β-carotene accelerated by sulfides under UVA irradiation were also identified. The results indicated that the disulfides had more obvious accelerative effects on the photodegradation of β-carotene than mono sulfides. The degradation of β-carotene after methyl (2-methyl-3-furyl) disulfide (MMFDS), methyl furfuryl disulfide (MFDS) and bis(2-methyl-3-furyl) disulfide (BMFDS) exposure followed first-order kinetics. Furan-containing sulfides such as MMFDS and BMFDS showed more pronounced accelerative effects than their corresponding isomers. The oxidation products were identified as 13-*cis*-β-carotene, 9,13-di-*cis*-β-carotene and all-*trans*-5,6-epoxy-β-carotene. These results suggest that both the sulfur atom numbers and the furan group in oxygen-containing sulfides play a critical role in the photooxidation of β-carotene.

**Keywords:** oxygen-containing sulfur flavor molecules; β-carotene; bis(2-methyl-3-furyl) disulfide (BMFDS); oxidation products

#### **1. Introduction**

Carotenoids are natural pigments of the isoprenoid family, commonly biosynthesized in fruits and vegetables [1], presenting potential physiological benefits, such as antioxidants in food and pro-vitamin A activity [2]. As one of the most commonly used carotenoids, β-carotene is expected to be conducive to health because of its valuable nutritional properties and antioxidant capacities, which confer on this compound an important role in lowering the risk of cataracts [3], inhibiting age-related macular degeneration [4], and enhancing the prevention of cardiovascular diseases [5].

However, due to its poor water solubility [6] and low bioavailability [7,8] during food processing and storage, widespread applications of β-carotene in food matrices normally suffer considerable challenges. Moreover, the restriction of β-carotene utilization as a nutritional ingredient in the food industry is currently also attributed to the existence of numerous unsaturated groups, resulting in high vulnerability to degradation reaction when exposed to light, heat and other external factors [9,10]. It has been reported that strong illumination can influence the stability of β-carotene extracted from palm oil, unveiling the formation of cis isomers [11]. The in-depth study carried out by Ayu et al. [12], who investigated interactive influence of tocopherols, tocotrienols, and β-carotene in the process of

photooxidation of red palm oil, suggesting that the degradation of β-carotene easily occurs under the irradiation of light. Apart from that, a comparison of β-carotene degradation under different UV stresses was conducted by Chen et al. [13], which showed that the longer the wavelength applied, the faster the degradation rate.

Several reports have also focused on the effects of chemical substances and their stability. The presence of 1,4-dimethylnaphthalene-1,4-endoperoxide and lycopene had the potential to induce the generation of (9*Z*)-, (13*Z*)- and (15*Z*)-β-carotene, which was associated with the formation of singlet oxygen [14]. Lewis acids, including titanium tetrachloride and ferric chloride, can catalyze the degradation of β-carotene to form an intermediate radical carbocation [15].

Currently, more than 300 sulfides have been registered as Generally Recognized as Safe (GRAS) substances with various threshold limits, making them the critical food flavors [16]. Biological functions including antithrombotic [17], antimicrobial [18], anticancer [19], and anti-inflammatory activities [20] in combination with their attractive odor characteristics such as garlic, onion, meat and nut flavors, have increased their feasibility of acting as food additives. In our previous studies, we discovered that dimethyl sulfides exerted apoptosis-inducing effects in leukemia cell lines via the generation of reactive oxygen species, especially for dimethyl trisulfide(Me2S3) and dimethyl tetrasulfide(Me2S4) [21]. Furthermore, β-carotene combined with Me2S<sup>4</sup> under UVA irradiation presented a synergistic action in inhibiting the viability of HL-60 cells viability, and elevating caspase-3 levels [22], mostly like probably raising the possibility of the reaction between sulfides and β-carotene assisted by UVA.

In this study, we selected 11 kinds of oxygen-containing sulfur flavor molecules, commonly used in the food industry, as experimental materials to examine their influence on β-carotene stability under UVA irradiation. Moreover, both the dynamic analysis of β-carotene degradation and the structural effects of sulfides that accelerated the degradation of β-carotene were investigated to provide a clearer and better comprehension of their acceleration effects. Furthermore, the oxidation products of β-carotene under UVA irradiation were also analyzed in order to elucidate its degradation mechanism.

#### **2. Results**

#### *2.1. The Effects of Oxygen-Containing Sulfur Flavor Molecules on β-Carotene Degradation under UVA Irradiation*

The structures of 11 kinds of oxygen-containing sulfur flavor molecules are shown in Table 1. Most of the sulfides contain furan or furfuryl group, such as 2-methyl-3-(methylthio) furan (MMTF) and methyl furfuryl disulfide (MFDS). Some sulfides contain different numbers of sulfur atoms but have the same side chain groups, such as MMTF and methyl (2-methyl-3-furyl) disulfide (MMFDS), difurfuryl sulfide (DFS) and difurfuryl disulfide (DFDS). Furthermore, it is worth noting the existence of isomers, such as bis(2-methyl-3-furyl) disulfide (BMFDS) and DFDS, MMFDS and MFDS.

The effects of 11 kinds of oxygen-containing sulfur flavor molecules assisted by UVA irradiation on β-carotene stability are shown in Figure 1A. After irradiating under UVA for 60 min in which the light intensity was 2.5 mW/cm<sup>2</sup> , the contents of β-carotene in all groups showed a reducing trend. To gain more knowledge about the correlation between the structure of coexistent sulfides and the degradation ratios of β-carotene, the remaining amounts of β-carotene were compared according to the structural characteristics of coexistent sulfides. The amounts of β-carotene treated with MMFDS and BMFDS were dramatically decreased by approximately 96.05% and 99.70%, respectively (*p* < 0.05). Likewise, the remaining amount of β-carotene in the presence of MFDS declined approximately 43.64%. These findings proved the fact that natural sulfur substance may affect β-carotene stability.

#### *2.2. The Effects of Furan-Containing Sulfides on β-Carotene Degradation under UVA Irradiation*

The order of the reaction with respect to the photodegradation of β-carotene was acquired according to Equation (2) to examine the changes of β-carotene concentration with time after UVA irradiation.


**Table 1.** Structure of eleven oxygen-containing sulfur flavor molecules.

β

β

β

S S

β

O

O

FEMA: Flavor and Extract Manufacturers Association of the United States. O

β β β β **Figure 1.** The effect of oxygen-containing sulfides on the degradation of β-carotene in ethanol under UVA irradiation. β-Carotene was treated with various oxygen-containing sulfur flavor molecules under UVA irradiation for 60 min (**A**) or within 60 min (**B**) in ethanol system. The residual β-carotene (**A**) and the first-order kinetics curve of β-carotene degradation (**B**) were determined. The bar results are expressed as means ± SD from three independent replicates. Different small letters show significant differences (*p* < 0.05).

As presented in Figure 1B, β-carotene degradation upon exposure to MMFDS, BMFDS and MFDS followed first-order kinetics, consistent with the kinetic model in dichloromethane system [23]. The corresponding kinetic parameters are listed in Table 2. It can be seen that the presence of BMFDS in ethanol significantly improved the k and shortened the t1/2 of β-carotene degradation, compared with the other two sulfides. The k and t1/2 were 0.131 min−<sup>1</sup> and 5.29 min for BMFDS, while they were 0.0633 min−<sup>1</sup> and 10.95 min for MMFDS and 0.0095 min−<sup>1</sup> and 72.96 min for MFDS, respectively. Furthermore, it should also be noticed that these three kinds of sulfides (MMFDS, BMFDS and MFDS) having obviously promoting effects on the degradation of β-carotene contain at least two sulfur atoms, which is consistent with the previous report [24]. These results suggest that sulfides with more sulfur atoms might trigger a stronger chemical effect on β-carotene stability in ethanol model system.

**Table 2.** Degradation kinetics parameters of β-carotene in the presence of BMFDS, MMFDS, MFDS under UVA irradiation in ethanol system.


BMFDS: Bis (2-methyl-3-furyl) disulfide, MMFDS: Methyl (2-Methyl-3-furyl) disulfide, MFDS: Methyl furfuryl disulfide.

#### *2.3. Kinetics of β-Carotene Degradation Treated with BMFDS under UVA Irradiation*

To further clarify the acceleration effect of BMFDS, the degradation kinetics parameters of β-carotene under UVA irradiation were determined. As shown in Table 3, the photodegradation of β-carotene treated by BMFDS in an ethanol system followed first-order kinetics. In terms of the rate constant k, the degradation degree of β-carotene treated with BMFDS was much higher, approximately 156 times than that in the control group. Therefore, our findings demonstrated that β-carotene degradation was followed first-order kinetics after treatment with BMFDS.

**Table 3.** Degradation kinetics parametersβ-carotene in the presence of BMFDS under UVA irradiation.


c is the concentration of reactant.

### *2.4. The Analysis of Photooxidation Products of β-Carotene Treated with BMFDS under UVA Irradiation*

According to the remarkable acceleration effect of BMFDS, HPLC-DAD-APCI-MS combined with Raman spectroscopy was applied to make a preliminary identification about the oxidation products of β-carotene, given their low contents and rather complex process of products collection. The chromatographic and spectral data of the oxidation products of β-carotene by HPLC-DAD-ACPI-MS are shown in Figure 2 and Table 4. The chromatographic peaks of leading β-carotene oxidation products had the same peak time in both the experimental group and the control group. Therefore, it was supposed that β-carotene treated with or without BMFDS under UVA irradiation had the same oxidation products (Figure 2A). In comparison to the retention time of a standard product (24.322 min) (Table 4), peak 3 was confirmed as all-*trans*-β-carotene (24.319 min) (Figure 3A). The UV spectra data are depicted in Figure 2B, and among the four obvious peaks, peak 2 and peak 4 were identified tentatively as 13-*cis*-β-carotene [25] and 9,13-di-*cis*-β-carotene [26], respectively (Figure 3C,D) according to the spectral characteristics and Q-ratios (Table 4), which were stipulated as the absorbance ratio of the middle main absorption peak to the cis peak.

**Table 4.** Product identification of β-carotene treated with BMFDS under UVA irradiation.


β β

β β **Figure 2.** The products of β-carotene treated with BMFDS under UVA irradiation. HPLC chromatograms (**A**) and UV-Vis spectra (**B**) of β-carotene oxidation products induced by BMFDS treatment for 5 min in ethanol under UVA irradiation. The blue line in HPLC chromatograms is referred to standard β-carotene. Peak identification for (**A**) and (**B**): Peak 1, all-*trans*-5,6-expoxy-β-carotene; Peak 2, 13-*cis*-β-carotene; Peak 3, all-*trans*-β-carotene; Peak 4, 9,13-di-*cis*-β-carotene. Raman spectra (**C**) at different wave number corresponded to β-carotene under UVA irradiation for 5 min with BMFDS (**a**) and without BMFDS (**b**), respectively.

β β

**λ**

β β β β

β

β

β β β

β β β β β **Figure 3.** The structure of β-carotene and its oxidation products induced by BMFDS under UVA irradiation. (**A**) all-*trans*-β-carotene; (**B**) all-*trans*-5,6-epoxy-β-carotene; (**C**) 13-*cis*-β-carotene; (**D**) 9,13-di-*cis*-β-carotene.

<sup>−</sup> β υ <sup>−</sup> υ <sup>−</sup> The Raman spectra at various wavenumber (cm−<sup>1</sup> ) of β-carotene in ethanol system treated with or without BMFDS under UVA irradiation are displayed in Figure 2C. By comparison of Figure 2C (a) and Figure 2C (b), both υ1 (–C=C–) at 1520.67 cm−<sup>1</sup> and υ2 (–C=C–C=C–) at 1155.48 cm−<sup>1</sup> can be observed. However, the distinction existed in the appearance of a new polar function group (C–O) presented in Figure 2C (a). Combined with its *m*/*z* of 553 (Table 4), this new vibration peak was identified tentatively as all-*trans*-5,6-epoxy-β-carotene (Figure 3B). As would have been expected, mono-epoxide is susceptible to generate from different sorts of carotenoids.

#### **3. Discussion**

β-Carotene was susceptible to be affected when exposed to external factors, consistent with previous reports, which considered that light can exert influence on β-carotene degradation [27–29]. In this study, sulfides have also showed to be involved in interfering the stability of β-carotene, which agreed with the report of Wei et al., implying that the stability of β-carotene can be significantly influenced when chitosan-(−)-epigallocatechin-3-gallate conjugates on β-carotene emulsions covered by sodium caseinate [30]. Moreover, in agreement with previous studies, both the number of sulfur atoms and the type of side group can affect the accelerated degradation of β-carotene under UVA irradiation [24,31]. It has been reported that the coexistence of disulfides can remarkably decrease the residual ratios of β-carotene to approximately 51.8–69.1%, while the presence of mono sulfides did not show obvious accelerating effects compared to the absence of mono sulfides [24].

β-Carotene degradation upon exposure to MMFDS, BMFDS and MFDS followed first-order kinetics, consistent with the kinetic model in dichloromethane system [23]. On the basis of the previous report which focused on the phenomenon of the existence of isomers [32], we also studied the accelerated effects of side groups among these sulfides on the degradation of β-carotene. Although MMFDS and MFDS both possess the same molecular formula (C6H8OS2), MMFDS showed a stronger accelerated degradation effect than MFDS. It is presumably because there is a methyl group and a furan group on the side of the disulfide bond in MMFDS, while a methyl group and a furfuryl group exist on the side of the disulfide bond in MFDS. For DFDS and its corresponding isomer BMFDS. Similarly, there is a furan group on both ends of the disulfide bond in BMFDS, while there is a furfuryl group on each side of the disulfide bond in DFDS. Their different abilities to promote the degradation of β-carotene can be related to the existence of various side groups. These results may account for the fact that furan-containing sulfur flavor molecules (MMFDS and BMFDS) showed a much more remarkable acceleration effect on the degradation of β-carotene than furfuryl-containing sulfur flavor molecules (MFDS and DFDS, respectively). Therefore, the number of sulfur atoms and the furan group in oxygen-containing sulfur flavor molecules may play a critical role in the accelerated degradation of β-carotene under UVA irradiation in ethanol system.

Several studies have investigated the order of kinetics on the degradation of β-carotene in different model systems under various conditions. It also followed a first order reaction under ambient storage, ultraviolet radiation and even heat treatments [13]. The photodegradation of β-carotene treated by BMFDS in ethanol system followed first-order kinetics, which agreed with previous studies carried out in food model systems such as carrots [33], oil/carrot emulsion system [34], oil model systems [10,35] and pulp or juices [36,37]. Ferreira et al. observed a first-order reaction for β-carotene degradation in a low-moisture and aqueous model system, as well as in lyophilized guava under different processing and storage conditions [38].

In addition, our results were in good agreement with Li et al. who summarized that the *trans*-*cis* isomerization of carotenoids can be generated via contacting with acids, thermal treatment or light [39]. It has been a long time since 13-*cis*-β-carotene was recognized as one of the main *cis* forms of β-carotene in food [40]. Chen et al. even analyzed it by different processing means, including over-heating and (non)-iodine-catalyzed photodegradation [41]. In addition, 9,13-di-*cis*-β-carotene was also confirmed as a common β-carotene degradation product according to Glaser et al. [42]. Moreover, our founding was consistent with Handelman et al., who had detected 5,6-epoxide of β-carotene through utilizing HPLC with mass analysis [43]. Similarly, Zeb identified all-*trans*-5,6-epoxy-β-carotene by an HPLC system and single ion monitoring mass spectrometry as well [44].

#### **4. Materials and Methods**

#### *4.1. Materials and Chemicals*

Eleven kinds of oxygen-containing sulfur flavors and β-carotene were obtained from Sigma-Aldrich (St. Louis, MO, USA). The structures of these sulfides are presented in Table 1. Methanol and methyl *tert*-butyl ether (MTBE)were procured from Damao Chemical factory (Tianjin, China) and Fisher Scientific (Pittsburgh, PA, USA), respectively. The other chemicals and reagents were of analytical grade.

#### *4.2. Preparation of the Model Systems*

For the preparation of the ethanol model system, 1 mg β-carotene was dissolved in 15 mL ethanol according to Onsekizoglu et al. [27] with minor modifications. The β-carotene solution was prepared daily and kept in the dark at 4 ◦C before use. Stock solutions of 11 kinds of sulfides were prepared in ethanol at a concentration of 10 mM and kept at 4 ◦C prior to use.

#### *4.3. Kinetic Analysis of β-Carotene Degradation*

The working solutions of β-carotene were transferred into quartz cuvettes, followed by the addition of 10 µL sulfur flavors. The control was performed with 10 µL ethanol. Then, the mixture was treated by UVA light (2.5 mW/cm<sup>2</sup> ) with the aim of assessing the degradation kinetics of β-carotene treated with sulfide. The degradation of β-carotene was measured immediately in a UV-1750 spectrophotometer (Shimadzu, Tokyo, Japan) at the wavelength of 450 nm for 60 min, which was monitored every 10 min. All measurements were performed in triplicate and data are expressed as mean of three independent experiments.

#### *4.4. Degradation Kinetics Modeling*

The trial-and-error procedure was carried out in accordance with the integral method outlined by Sánchezet al. [45] to determine the reaction order of theβ-carotene degradation. Different order models can be represented as follows:

$$c - c\_0 = -kt\tag{1}$$

$$
\ln c/c\_0 = -kt\tag{2}
$$

$$1/c - 1/c\_0 = kt \tag{3}$$

In these formulas, *c* (µM) is thereactantconcentrationat a given time, *c*<sup>0</sup> (µM) is the initial reactant concentration, *k* (min−<sup>1</sup> ) is the degradation rate constant, and *t* (min) is the treatment time.

#### *4.5. Analysis of β-Carotene Treated with UVA Irradiation and BMFDS*

The working solutions of β-carotene were transferred into quartz cuvettes, followed by the addition of 10 µL BMFDS. The control was performed with 10 µL ethanol. Then, the mixture containing β-carotene and BMFDS was placed under a UV lamp (Shimadzu, Japan) with an intensity of 2.5 mW/cm<sup>2</sup> for 5 min, followed by drying completely under a nitrogen stream. The residue was redissolved in 0.1 mL MTBE before use.

The further analysis was carried out and relative parameters were applied according to Santos et al. [46]. Briefly, once redissolved, the solution was passed through a 0.22 µm filter, followed by the injection into an HPLC-DAD-APCI-MS system (Agilent, Santa Clara, CA, USA) for closer analysis. A YMC C<sup>30</sup> column (250 × 4.6 mm, 5 µm) and gradient mobile phase of methanol-MTBE-water (85:15:5, *v*/*v*/*v*) and MTBE (100%) were used for β-carotene detection.

#### *4.6. Determination of Degradation Products by Raman Spectroscopy*

The Raman spectra of the degradation products were recorded on a Raman spectrometer (Bruker Instruments Inc., Bill-erica, MA, USA). The wave number was in the range of 400–4000 cm−<sup>1</sup> using the 785 nm as the excitation line. The power was 10 Mw while the integration time was 20 s.

#### *4.7. Statistical Analysis*

All the data were expressed as the mean ± SD or mean and subjected to the Student's *t*-test for statistical analysis. Statistical significance was considered at a *p* < 0.05.

#### **5. Conclusions**

Overall, through applying the results we obtained and drawing upon the information provided by other studies, it is supposed that the oxidation products of β-carotene treated with BMFDS under UVA irradiation in ethanol system might include 13-*cis*-β-carotene, 9,13-di-*cis*-β-carotene and all-*trans*-5,6-epoxy-β-carotene. Our results might shed new light on the accelerative effect of BMFDS on the photodegradation of β-carotene. More insights into the mechanism involved in degradation of oxidation products of BMFDS-treated β-carotene should be further studied further.

**Author Contributions:** Methodology, Y.L. and W.-Q.G. formal analysis, J.S. data curation, Y.L. and J.S.; writing—original draft preparation, G.-L.Z. and H.-Y.W.; writing—review and editing, G.-L.Z. and H.-M.H.; supervision, H.-M.H.; funding acquisition, G.-L.Z. All authors discussed the results and approved the final manuscript.

**Funding:** This research was funded by The National Natural Science Foundation of China, grant numbers [31571888 and 31201419].

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

#### **References**


**Sample Availability:** Samples of the compounds are not available from the authors.

© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*
