Journal of Food Quality

Journal of Food Quality / 2017 / Article

Research Article | Open Access

Volume 2017 |Article ID 8401074 |

Adriela Albino Rydlewski, Damila Rodrigues de Morais, Eliza Mariane Rotta, Thiago Claus, Julianna Matias Vagula, Michele Cristina da Silva, Oscar Oliveira Santos Junior, Jesuí Vergílio Visentainer, "Bioactive Compounds, Antioxidant Capacity, and Fatty Acids in Different Parts of Four Unexplored Fruits", Journal of Food Quality, vol. 2017, Article ID 8401074, 9 pages, 2017.

Bioactive Compounds, Antioxidant Capacity, and Fatty Acids in Different Parts of Four Unexplored Fruits

Academic Editor: Ángel A. Carbonell-Barrachina
Received17 Feb 2017
Revised20 Mar 2017
Accepted06 Apr 2017
Published24 Apr 2017


Total phenolic content (TPC), total flavonoids (TF), total anthocyanins (TA), and antioxidant capacity of different parts of four unexplored fruits from Brazil (Syzygium cumini Lam, Solanum nigrum Linn, Inga edulis Mart, and Hovenia dulcis Thunb) were determined; the bioactive compounds and fatty acids were quantified and identified by high-performance liquid chromatography and gas chromatography, respectively. S. cumini peels contained the most TA (63.31 mg/100 g), whilst H. dulcis pulp and peels had the highest TPC (518.18 mg GAE/100 g) and TF (76.54 mg EQ/g). Phenolic compounds responsible for antioxidant capacity of fruits were gallic acid, ellagic acid, kaempferol, and epicatechin. H. dulcis seed showed the highest level of the essential fatty acid omega-3 (3985.95 mg/100 g). PCA showed that PC1 and PC2 explained 90.43% of the total variability of the antioxidant data. Most of the seeds showed omega-3, omega-6, and omega-9 fatty acids at significant concentrations, with two PCs explaining 93.80% of the total variance of the fatty acid contents.

1. Introduction

Oxidative stress is caused by reactive oxygen species (ROS), which include free radicals (oxygen and nitrogen species), and this condition occurs when there is an overproduction of ROS and/or insufficient antioxidant mechanisms [1]. Among the main consequences of oxidative stress is damage to the biomolecules, particularly DNA, lipids, and proteins, which can severely compromise cell function and activate a cascade of free radical reactions. Oxidative stress may occur as a result of degenerative diseases or may also be a causative factor in the development of diseases. Arthritis, atherosclerosis, cancer, diabetes, and Parkinson’s, for example, involve ROS [2].

The interest in studying unexplored species of fruits and their parts, such as peels and seeds, has increased in recent years because they are rich in phenolic compounds, flavonoids, anthocyanins, vitamins, minerals, and other bioactive compounds that are capable of neutralizing free radicals [35]. Moreover, fruits have essential polyunsaturated fatty acids (PUFAs), such as linoleic acid (LA, 18:2n-6) and alpha-linolenic acid (LNA, 18:3n-3) that are considered essential because they cannot be synthesized by the human body and must be supplied through the diet [6]. These fatty acids (FAs) are able to protect against oxidative stress by exerting an antioxidant role [7]. Fruits also contain oleic acid (OA, 18:1n-9), which is widely recognized for its action in lowering the risk of ROS-induced diseases and enhancing the immune system [8]. Therefore, increased fruit consumption is associated not only with personal preference but also with the maintenance of health, through the consumption of foods rich in vital micronutrients [7, 9].

Several methods have been developed to measure the antioxidant capacity of fruits. In particular, the ferric reducing antioxidant power (FRAP) and 1,1-diphenyl-2-picrylhydrazyl (DPPH) assays have been widely adopted [7]. Association studies between fruit and vegetable consumption and indicators of oxidative stress using antioxidant assays have received the attention of researchers [10]. It is recommended that at least two methods with different mechanisms are combined to provide a reliable picture of the total antioxidant capacity of a food because the results can be affected using a single assay to evaluate the entire sample [11, 12].

Jambolan (Syzygium cumini Lam), black nightshade (Solanum nigrum Linn), inga edulis (Inga edulis Mart), and Japanese grape (Hovenia dulcis Thunb) are unexplored fruits and few studies have been published on characterizing the bioactive compounds and antioxidant capacity of the pulps, peels, and seeds of these fruits. Therefore, this study aimed to evaluate the DPPH and FRAP antioxidant capacities, the total phenolic content (TPC), total flavonoids (TF), and total anthocyanins (TA) and to identify and quantify the phenolic compounds and FAs by high-performance liquid chromatography (HPLC) and gas chromatography (GC), respectively.

2. Materials and Methods

2.1. Plant Material

The botanical identification and geographical origin of the four fruits are provided in Table 1. Fresh fruits (ca. 2 kg each) were harvested at Monte Alegre, São Paulo, Brazil (23°35′31′′S and 48°38′38′′W). The fruits were washed with tap water and the pulps, peels, and seeds were removed manually, chopped, and then homogenized in a multiprocessor to uniformity before analysis. H. dulcis pulp and peels were evaluated together because it was not possible to separate them. I. edulis peels were discarded.

Common nameScientific nameFamilyOrigin

JambolanSyzygium cumini LamMyrtaceaeIndia
Black nightshadeSolanum nigrum LinnSolanaceaeSouth Africa
Inga edulisInga edulis MartLeguminosaeBrazil, South and Central America
Japanese grapeHovenia dulcis ThunbRhamnaceaeJapan, China, and Korea

2.2. Chemical and Standards

Fatty acid methyl esters (FAME) (standard mixture 189-19), methyl tricosanoate (23:0), Folin-Ciocalteu phenol reagent, gallic acid, kaempferol, ellagic acid, epicatechin, quercetin, DPPH, 6-hydroxy-2,5,7,8-tetramethylchromane-2-carboxylic acid (Trolox), 2,4,6-tris (2-pyridyl)-s-triazine (TPTZ), and methanol and acetonitrile (HPLC grade) were purchased from Sigma Chemical Co. (St. Louis, USA). Ultrapure water (Milli-Q system, Millipore Corp, Bedford, MA) was also used. All solvents and chemicals were of analytical grade.

2.3. Extraction of Antioxidants

The extracts were prepared using 10 g (per part of the fruit) in 100 mL of methanol under magnetic stirring for 4 h according to the method developed by Santos et al. [13]. After filtration, the extracts were concentrated under reduced pressure at 40°C. The absorbance values obtained were within the calibration range of the respective methods (Genesys 10-UV scanning, Thermo Scientific). The results were expressed as fresh weight (FW).

2.4. Analysis of DPPH Antioxidant Capacity (Free Radical-Scavenging Activity)

The DPPH scavenging capacity was measured using the method described by Brand-Williams et al. [14]. Briefly, an aliquot of the methanolic fruit extract (25 µL) was added to 2 mL of 6.25 × 10−5 mol/L DPPH methanolic solution. After gently mixing and then resting the reaction solutions at room temperature for 30 min, the absorbance was measured at 517 nm. Methanolic solutions of known Trolox concentrations between 0 and 2000 µmol/L were used for calibration. The results were expressed as µmol Trolox equivalents (TE)/g FW using the calibration curve (, ).

2.5. FRAP Assay

The FRAP assay was determined as previously described by Benzie and Strain [15]. FRAP reagent was prepared by mixing acetate buffer (0.3 mol/L, pH 3.6), TPTZ (10 mmol/L), and FeCl3 (20 mmol/L) solutions at 10 : 1 : 1, respectively. An aliquot of the methanolic fruit extract (100 µL) and 300 µL of distilled water were added to 3 mL of the FRAP reagent and then placed in the dark for 30 min at 37°C. The absorbance was measured against a blank at 593 nm. Aqueous solutions of known Fe2+ concentrations between 0 and 1500 µmol/L (FeSO4·7H2O) were used for the calibration curve (, r2 = 0.999) and the results were expressed as µmol Fe2+/g FW.

2.6. TPC

The TPC were determined according to the procedure described by Shahidi and Naczk [16]. An aliquot of the methanolic fruit extract (250 μL) was mixed with 250 μL of the Folin-Ciocalteu reagent (prediluted in distilled water at 1 : 1 v/v), 500 μL of a saturated sodium carbonate solution, and 4 mL of distilled water. The reaction mixture was left to rest for 25 min and then centrifuged at 3000 rpm (1638 g) for 10 min. Then, the absorbance was measured at 725 nm. Methanolic solutions of known gallic acid concentrations between 0 and 250 mg/L were used for calibration. The results were expressed as mg gallic acid equivalents (GAE) 100/g FW using the calibration curve (, r2 = 0.999).

2.7. TF

TF were determined according to Woisky and Salatino [17]. An aliquot of the methanolic fruit extract (500 µL, 2.5 mg/ml) was added to 250 µL of AlCl3 (5 g/100 mL methanol) and 4.25 mL of methanol. After 30 min, the absorbance was measured at 425 nm. The results were expressed as mg quercetin equivalents (mg QE/g FW) using the calibration curve (, r2 = 0.999).

2.8. TA

TA were extracted according to Lees and Francis [18]. An aliquot (50 g) was then homogenized with 50 mL of solvent (70% ethanol acidified to pH 2.0 with 0.1% HCl) for 2 min. The volume was adjusted to 200 mL in a volumetric flask using the solvent, covered with parafilm, and stored at 4°C for 12 h. Then, the material was filtered using a Büchner funnel and 125 mL of the filtrate mixed with another 250 mL of the solvent. Triplicate aliquots (2 mL) were adjusted to a final volume of 100 mL in a volumetric flask using the solvent. The reaction mixture was placed in the dark for 2 h. Then, the absorbance was measured at 535 mm. The results were expressed as mg 100/g FW.

2.9. Chromatographic Analysis

Total lipids (TL) were extracted by Bligh and Dyer [19] method. FAME were prepared by methylation of the TL as described by Hartman and Lago [20] and the methyl esters separated by GC in a Trace Ultra model 3300 (Thermo Scientific) equipped with a flame ionization detector (FID) and a cyanopropyl capillary column (100 m × 0.25 i.d., 0.25 μm film thickness, CP7420). The gas flow rates were 1.2 Ml/min carrier gas (H2), 30 mL/min make-up gas (N2), and 35 and 300 mL/min flame gasses (H2 and synthetic air, respectively). The split ratio was 1 : 80 and 2 μL of the samples was injected in triplicate. The detector and injection port were set at 240°C and the column was set at 185°C for 7.5 min. The peak areas were determined by ChromQuest 5.0 software. The FAs were identified by comparing their retention times with those of standard methyl esters.

Quantification (as mg FA/g of TL) was determined against tricosanoic acid methyl ester (23:0) as an internal standard (IS), as described by Joseph and Ackman [21]. Theoretical FID correction factor [22] values were used to obtain the concentrations. The FA contents were calculated as mg/g TL using where FA is expressed as mg/g of TL, is the peak area (FA), is the peak area of the IS (the methyl ester of tricosanoic acid, 23:0), is the IS mass (mg) of IS added to the sample (mg), is the sample mass (mg), is the theoretical correction factor, and is the conversion factor necessary to express results as mg of FA rather than as methyl esters.

2.10. Quantification of Phenolic Compounds by HPLC

The phenolic compounds were quantified according to Fratianni et al. [23] using a Thermo Scientific chromatograph, model Finnigan Surveyor PDA, with a manual injection valve, a photodiode array detection (DAD), connected to Crom-Quest 5.0 software and equipped with a EC 250/4.6 Nucleodur 100-5 C18ec column (Macherey-Nagel) (250 mm × 4.6 mm). The mobile phases consisted of Milli-Q water (containing 1% acetic acid, phase A) and acetonitrile (containing 1% acetic acid, phase B). The gradient started at 90% A and decreased to 60% A in 25 min. After 26 min, phase A returned to 90%, where it remained until 35 min (total run time). The flow rate was 0.7 ml/min, the injection volume was 50 µl, and the detection wavelengths were set at 250 and 270 nm. The phenolic compounds of the methanolic fruit extracts were identified by comparison of their retention times with standards (Sigma) and UV-visible light spectra. The bioactive compounds were quantified by comparing the sample with the standard calibration curves.

2.11. Statistical Analysis

The data were analyzed using analysis of variance (ANOVA) and the means were compared by Tukey’s test. The results were analyzed by principal component analysis (PCA) using Statistica 7.0 software. Data before treatment was not necessary.

3. Results and Discussion

3.1. TPC, TF, TA, and Antioxidant Capacity

The TPC, TF, and TA amounts and the antioxidant capacity (DPPH and FRAP) of the different parts of the fruits are shown in Table 2. The TPC ranged from 19.27 mg GAE/100 g in S. nigrum seeds to 518.18 mg GAE/100 g in H. dulcis pulp and peels. The TPC in H. dulcis seed (85.56 mg GAE/100 g) was similar to that found by Santos et al. [13] in Benitaka and Brazil grape seed varieties (89.83 and 91.53 mg GAE/100 g, respectively). The TPC of S. nigrum pulps was 412.71 mg GAE/100 g. This value can be compared to those of jaboticaba (440 mg GAE/100 g), açaí (454 mg GAE/100 g), and gurguri (549 mg GAE/100 g), in the study by Rufino et al. [9] because of the scarcity of studies on this particular fruit.

Parts of fruitsAnalysis

Syzygium cumini Lam ± 8.93 ± 2.36 ± 0.41 ± 114.56 ± 10.30
Solanum nigrum Linn ± 9.46 ± 5.95 ± 1.09 ± 105.83 ± 12.54
Inga edulis Mart ± 7.36 ± 1.01 ± 0.05 ± 9.54 ± 3.61
Hovenia dulcis Thunb ± 30.89 ± 5.47 ± 1.14 ± 309.61 ± 9.43
Syzygium cumini Lam ± 7.84 ± 7.34 ± 1.64 ± 32.49 ± 6.08
Solanum nigrum Linn ± 1.38 ± 5.44 ± 0.12 ± 1.27 ± 3.38
Inga edulis MartNENENENENE
Hovenia dulcis Thunb ± 30.89 ± 5.47 ± 1.14 ± 309.61 ± 9.43
Syzygium cumini Lam ± 9.46 ± 3.37 ± 0.18 ± 639.45 ± 3.77
Solanum nigrum Linn ± 1.06 ± 1.42 ± 0.28 ± 1.15 ± 1.11
Inga edulis Mart ± 8.68 ± 0.69ND ± 12.43 ± 3.46
Hovenia dulcis Thunb ± 7.28 ± 4.36 ± 0.25 ± 195.27 ± 1.22

Results expressed as mean ± standard deviation of three replicates. Equivalent parts of the different fruits were compared. Means followed by different letters in the same column are significantly different by Tukey’s test ( < 0.05). H. dulcis pulp and peel were evaluated together. ND = not detected. NE = not evaluated.

TF among the parts of the fruits varied from 2.59 mg EQ/g in I. edulis seeds to 76.54 mg EQ/g in H. dulcis pulps and peels. According to Volp et al. [24], flavonoids are polyphenolic structures found naturally in plants and many of these flavonoids are responsible for the color of fruits. In the current study, H. dulcis pulp and peels revealed a higher TF value compared to the other pulps evaluated, which could be associated with the heavy pigmentation of these fruits.

Faria et al. [25] detected 91.2 mg EQ/g TF in S. cumini. These values were higher than those found in this study (59 mg and 65.51 mg EQ/g in pulps and peels, respectively). In contrast, Benherlal and Arumughan [26] studied the same fruit and only found 7 mg EQ/g. These variations may occur due to the influence of various factors, such as climate conditions, geographic location, fruit maturation stage, and method used [25, 27].

The highest TA concentration was in S. cumini peels (63.31 mg/100 g). This value was lower than that found by Rufino et al. [9], when evaluating the combined peel and pulp of the fruit (93.3 mg/100 g) but higher when compared with jaboticaba (58.1 mg/100 g) and camu-camu (42.2 mg/100 g), in the same study. Relatively low TA concentrations were found in I. edulis pulps (0.06 mg/100 g) and none were found in the seeds of this fruit. The DPPH antioxidant capacity was lower in the pulp of this fruit, compared to the other pulps evaluated in this study, in agreement with its lower content of bioactive compounds.

The DPPH antioxidant capacity of the fruit extracts varied from 68.66 μmol TE/g in S. nigrum peels to 13240.60 μmol TE/g in H. dulcis peel and pulp. The FRAP antioxidant activity varied from 22.93 µmol Fe2+/g in S. nigrum seed to 690.68 µmol Fe2+/g in H. dulcis pulp and peel. Both methods indicated that H. dulcis pulp and peel had the highest antioxidant capacity, probably due to having the highest TPC. Rufino et al. [9] evaluated the FRAP antioxidant capacity of methanolic extracts (methanol-water, 50 : 50 v/v) from the combined pulps and peels of S. cumini and found 35.5 µmol Fe2+/g. This value was lower than that found in the current study, in pulp and peel of the same fruit (341.95 and 323.50 µmol Fe2+/g, respectively).

3.2. Identification of Phenolic Compounds by HPLC

The phenolic compounds responsible for the antioxidant capacity of the methanolic fruit extracts were gallic acid, kaempferol, epicatechin, and ellagic acid (with peak retention times of 3.6, 5.4, 14.7, and 19.1 min, respectively) (Table 3). The flavonoid, kaempferol, showed at 250 nm and epicatechin at 270 nm. The gallic and ellagic acids also showed at 270 nm, characteristic of phenolic acids derived from hydroxybenzoic acid [25].

Parts of fruitsPhenolic compounds
Gallic acidKaempferolEpicatechinEllagic acid

Syzygium cumini Lam ± 0.09 ± 5.67NDND
Solanum nigrum Linn ± 0.08 ± 5.49NDND
Inga edulis MartND ± 6.34NDND
Hovenia dulcis Thunb ± 0.21 ± 7.84NDND
Syzygium cumini Lam ± 0.04 ± 7.98NDND
Solanum nigrum Linn ± 0.02 ± 12.46NDND
Inga edulis MartNENENENE
Hovenia dulcis Thunb ± 0.21 ± 7.84NDND
Syzygium cumini Lam ± 5.67 ± 5.634.81 ± 0.573.63 ± 4.13
Solanum nigrum Linn ± 0.04 ± 6.92NDND
Inga edulis Mart ± 0.46 ± 4.13NDND
Hovenia dulcis Thunb ± 9.84 ± 5.25NDND

Results expressed as mean ± standard deviation of three replicates. Equivalent parts of the different fruits were compared. Means followed by different letters in the same column are significantly different by Tukey’s test ( < 0.05). H. dulcis pulp and peel were evaluated together. ND = not detected. NE = not evaluated.

Gallic acid was the main contributor to the antioxidant capacity of S. cumini seeds (57.10 mg/100 g) and H. dulcis seeds (34.61 mg/100 g). S. cumini pulps presented 5.59 mg/100 g of gallic acid, which was comparable to that found in walnut (3.39 mg/100 g) by Donno et al. [28]. As stated previously, phenolic compounds, such as gallic acid, exhibit potent antioxidant, antidiabetic, and antihyperlipidemic activities [29].

Although the H. dulcis pulp and peel showed the highest TPC content and the highest DPPH and FRAP antioxidant capacities, it did not have the highest content of gallic acid. This was probably due to the limited interaction between the extraction solvent and gallic acid due to the high reactivity of this phenolic compound with oxidative materials and macromolecules. These factors make the identification process complex [30].

Kaempferol ranged from 11.45 mg/100 g in I. edulis pulps to 57.38 mg/100 g in S. nigrum peels. Using HPLC-DAD/mass spectrometry, Donno et al. [5] found 0.55 mg/100 g of kaempferol in strawberries, being a value lower than that found in all parts of fruits in this study. Ellagic acid and epicatechin were only identified in S. cumini seeds (3.63 and 4.81 mg/100 g, respectively). Epicatechin has been shown to exert several physiological effects, such as antioxidant activity and antitumor effects [31]. More recently, Rodríguez-Ramiro et al. [32] have shown that epicatechin was able to activate the detoxifying enzyme glutathione S-transferase pi 1 (GSTP1) related to the neutralization of reactive oxygen species and nitrogen species, in epithelial colon cells. Flavonols, like kaempferol and ellagic acid, possess health-promoting properties and may also contribute to the neutralization of free radicals and the maintenance of heart health [28, 33].

3.3. Fatty Acids (FAs)

A total of eight FAs were detected in pulps, peels, and seeds of fruits (Table 4). S. nigrum peels presented 1212.20 mg/100 g in the sum of saturated fatty acid (), representing 87% of TL. Among the seeds, I. edulis presented the highest values of SFA, followed by H. dulcis (613.47 mg/100 g and 526.66 mg/100 g, respectively), showing a significant difference between them.

Parts of fruitsFatty acids

Syzygium cumini Lam2.19c ± 0.036.38c ± 0.050.26b ± 0.0118.19b ± 0.175.10d ± 0.030.10d ± 0.010.04b ± 0.010.08c ± 0.018.61c ± 0.0818.52b ± 0.105.20d ± 0.030.60b ± 0.015.10d ± 0.030.10d ± 0.0151.01a ± 0.24
Solanum nigrum Linn1.05c ± 0.157.88c ± 0.278.63a ± 0.230.20d ± 0.0948.09b ± 6.020.94b ± 0.230.27a ± 0.380.10c ± 0.039.20c ± 0.748.93d ± 0.9349.03b ± 6.255.32a ± 0.2348.09b ± 6.020.94b ± 0.2351.15a ± 1.41
Inga edulis Mart11.66b ± 4.7413.93a ± 3.460.07c ± 0.0197.36a ± 33.88.45c ± 10.560.22c ± 0.010.05b ± 0.010.47a ± 0.0525.64b ± 2.9297.90a ± 12.398.67c ± 0.120.33c ± 0.058.45c ± 10.560.22c ± 0.0138.40b ± 5.68
Hovenia dulcis Thunb58.28a ± 9.958.10b ± 0.249.02a ± 0.202.14c ± 0.2398.69a ± 10.4741.36a ± 8.580.36a ± 0.020.25b ± 0.0166.74a ± 2.2611.41c ± 0.32140.05a ± 19.302.09a ± 0.1798.69a ± 10.4741.36a ± 8.582.38c ± 0.14
Syzygium cumini Lam29.66c ± 4.9283.33a ± 5.282.64c ± 0.78257.24a ± 27.0358.44a ± 3.552.99b ± 0.340.12b ± 0.071.46b ± 0.17112.11b ± 10.13261.34a ± 9.3361.43b ± 7.520.54b ± 0.1858.44a ± 3.552.99c ± 0.3419.54a ± 1.79
Solanum nigrum Linn299.55a ± 13.9224.43b ± 0.9934.41a ± 2.351.69b ± 0.021.46b ± 0.206.04b ± 0.82888.22a ± 43.50138.01a ± 32.201212.20a ± 56.44174.11b ± 12.197.50c ± 0.230.01c ± 0.011.46b ± 0.206.04b ± 0.820.24c ± 0.02
Hovenia dulcis Thunb58.28b ± 9.958.10c ± 0.249.02b ± 0.202.14b ± 0.2398.69a ± 10.4741.36a ± 8.580.36b ± 0.020.25b ± 0.0166.74c ± 2.2611.41c ± 0.32140.05a ± 19.302.09a ± 0.1798.69a ± 10.4741.36a ± 8.582.38b ± 0.14
Syzygium cumini Lam1.56c ± 0.450.95d ± 0.100.24d ± 0.12291.01c ± 41.700.45d ± 0.057.08d ± 0.160.40c ± 0.151.55c ± 0.432.91d ± 0.26292.79c ± 9.387.53d ± 0.212.58c ± 0.130.45d ± 0.057.08d ± 0.160.06d ± 0.01
Solanum nigrum Linn37.90b ± 5.3715.14c ± 1.062.67b ± 0.381891.36a ± 123.18957.72b ± 9.5012.23c ± 6.40116.47a ± 2.411.07c ± 0.08169.51c ± 96.711895.10a ± 176.61969.95b ± 14.685.72b ± 0.34957.72b ± 9.5012.23c ± 6.4078.30a ± 3.04
Inga edulis Mart313.70a ± 52.29299.17a ± 6.131.09c ± 0.04192.85d ± 9.989.84c ± 0.0460.76b ± 0.420.60c ± 0.049.41b ± 0.03613.47a ± 82.29203.35d ± 3.5770.60c ± 2.100.11d ± 0.019.84c ± 0.0460.76b ± 0.420.16c ± 0.01
Hovenia dulcis Thunb326.76a ± 34.29161.06b ± 7.8966.69a ± 7.56994.30b ± 24.541935.12a ± 120.173985.95a ± 652.2638.84b ± 4.9250.72a ± 2.75526.66b ± 5.821111.71b ± 2.285921.07a ± 786.2511.24a ± 1.221935.12a ± 120.173985.95a ± 652.260.48b ± 0.04

Results expressed as mean ± standard deviation of three replicates. Equivalent parts of the different fruits were compared. Means followed by different letters in the same column are significantly different by Tukey’s test (). H. dulcis pulp and peel were evaluated together. SFA: saturated fatty acid. MUFA: monounsaturated fatty acid. PUFA: polyunsaturated fatty acid. n-6: omega-6 fatty acid. n-3: omega-3 fatty acid. 16:0: palmitic acid. 18:0: stearic acid. 18:1n-7: cis-vaccenic acid. 18:1n-9: oleic acid. 18:2n-6: linoleic acid. 18:3n-3: α-linolenic acid. 20:0: arachidic acid. 20:1n-9: gondoic acid. NE = not evaluated.

S. nigrum seeds presented a value of 1895.10 mg/100 g in the sum of monounsaturated fatty acid (), representing 62.45% of TL. The major MUFA found in these seeds was OA (18:1n-9), showing a value of 1891.36 mg/100 g. This value of OA found was higher than those found by Santos et al. [13], in grape seed variety Isabel (1690.76 mg OA/100 g). The positive effect of dietary monounsaturated fats in preventing cancer has extensively been reported. In addition, the ingestion of this FA is associated with the improvement of the immune system due to its anti-inflammatory activity [8, 34].

H. dulcis seeds presented a value of 5921.07 mg/100 g in the sum of polyunsaturated fatty acid () (78.32% TL). The main PUFA was LNA (18:3n-3), which was found at the highest concentration in H. dulcis seeds (3985.95 mg/100 g). These seeds contained 8.73% TL, with 53% represented by LNA. Various n-3 PUFAs have been reported to exert beneficial effects, such as cholesterol regulation, blood pressure reduction, and cardiovascular disease prevention [35, 36].

According to Simopoulos [37], the relationship between PUFA/SFA should be greater than 0.4 to be adequate to prevent heart disease. The ratio of from all parts of the fruits has been higher than the recommended value, except in S. nigrum peels. This result was low because the peel of this fruit is rich in eicosanoic acid (20:0), presenting statistical difference among the other peels. This result also reflected the sum of saturated fatty acid () of the S. nigrum peels (1212.20 mg/100 g), which was the largest among all the peels.

The high n-6 to n-3 ratio in the diets of Western countries which occurred after the introduction of modern agriculture and animal farming techniques has, at least partly, contributed to the rise in cardiovascular disease. Studies indicate that the human diet can decrease the chances of developing several diseases, so the nutrition societies recommend a healthy diet containing n-6 to n-3 ratio of 5 : 1 [38, 39]. The n-6 to n-3 ratio of the fruits ranged within 2.38–51.01 in pulps, 0.24–19.54 in peels, and 0.06–78.30 in seeds. All parts of the H. dulcis and the S. nigrum peels, S. cumini seeds, and I. edulis seeds presented an ideal ratio of n-6 to n-3.

3.4. Principal Component Analysis (PCA)

PC1 (68.70%) and PC2 (21.73%) explained 90.43% of the total variance (Figure 1(a)). Loadings (variables)/score (samples) of antioxidant analyses of the fruits showed the formation of four groups (Figure 1(b)). One group consisted of DPPH, FRAP, TPC, TF, and sample 9, due to higher values in this sample, in all of these variables. In PC1, DPPH, FRAP, TPC, and TF contributed positively to the separation of this group, whilst, in PC2, DPPH, FRAP, and TPC contributed negatively and TF positively. Sample 2 was grouped with TA and also has highlighted the contents of this variable. Sample 3 showed high DPPH and TPC values; for this reason, its position was closer to these variables and separated from the other samples. A fourth group was formed by the remaining samples, which showed lower values in all analyses.

PC1 and PC2 accounted for 93.80% of the total variance of the FA data (Figure 2(a)) and 66.21% (PC1) of this variance was attributed to the positive contribution of the sum of PUFAs, n-6 and n-3 (Figure 2(b)). Sample 10 showed the highest values for these variables, forming group 1. The PUFAs and n-3 loadings of this group contributed negatively and only n-6 contributed positively to PC2, which accounted for 27.59% of the total variance. Sample 6 stood out in the MUFA values, contributing positively to PC2 and separating this group. The third group had the lowest values of the analyzed variables.

4. Conclusion

The bioactive compounds gallic acid and kaempferol were largely responsible for the antioxidant capacity of the parts of the fruits analyzed by HPLC, since the ellagic acid and the epicatechin were only found in S. cumini seeds. The parts of the fruits that stood out for their high TPC and TA contents showed a positive correlation between the contents of these compounds and the antioxidant capacity by the methods used. The PCA analysis showed that PC1 and PC2 explained 90.43% of the total variability of the antioxidant data. Most of the seeds studied showed omega-3, omega-6, and omega-9 FAs, in significant concentrations, with two PCs explaining 93.80% of the contribution of these FA contents and the sum of the MUFAs and PUFAs to these data.

Conflicts of Interest

The authors declare that there are no conflicts of interest regarding the publication of this paper.


The authors would like to thank the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior (CAPES) and the Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq) for their financial support and fellowships.


  1. B. Halliwell, “Free radicals and antioxidants: updating a personal view,” Nutrition Reviews, vol. 70, no. 5, pp. 257–265, 2012. View at: Publisher Site | Google Scholar
  2. R. L. Prior, “Oxygen radical absorbance capacity (ORAC): new horizons in relating dietary antioxidants/bioactives and health benefits,” Journal of Functional Foods, vol. 18, pp. 797–810, 2015. View at: Publisher Site | Google Scholar
  3. T. M. Rababah, N. S. Hettiarachy, and R. Horax, “Total phenolics and antioxidant activities of fenugreek, green tea, black tea, grape seed, ginger, rosemary, gotu kola, and ginkgo extracts, vitamin E, and butylhydroquinone,” Journal of Agricultural and Food Chemistry, vol. 52, no. 16, pp. 5183–5186, 2004. View at: Publisher Site | Google Scholar
  4. M. E. Schreckinger, J. Lotton, M. A. Lila, and E. G. De Mejia, “Berries from South America: a comprehensive review on chemistry, health potential, and commercialization,” Journal of Medicinal Food, vol. 13, no. 2, pp. 233–246, 2010. View at: Publisher Site | Google Scholar
  5. D. Donno, M. Cananna, G. L. Beccaro, M. G. Mellano, D. T. Marioni, and A. K. G. Bounous, “Currants and strawberries as bioactive compound sources: determination of antioxidant profiles with HPLC-DAD/MS system,” Journal of Applied Botany and Food Quality, vol. 86, pp. 1–10, 2013. View at: Google Scholar
  6. A. C. Aguiar, S. M. Cottica, M. Boroski et al., “Quantification of essential fatty acids in the heads of nile tilapia (Oreochromis niloticus) fed with linseed oil,” Journal of the Brazilian Chemical Society, vol. 22, no. 4, pp. 643–647, 2011. View at: Publisher Site | Google Scholar
  7. I. Gülçin, “Antioxidant activity of food constituents: an overview,” Archives of Toxicology, vol. 86, no. 3, pp. 345–391, 2012. View at: Publisher Site | Google Scholar
  8. S. Bilal, A. L. Khan, M. Waqas et al., “Biochemical constituents and in vitro antioxidant and anticholinesterase potential of seeds from Native Korean Persimmon Genotypes,” Molecules, vol. 21, no. 7, article 893, 2016. View at: Publisher Site | Google Scholar
  9. M. D. S. M. Rufino, R. E. Alves, E. S. de Brito, J. Pérez-Jiménez, F. Saura-Calixto, and J. Mancini-Filho, “Bioactive compounds and antioxidant capacities of 18 non-traditional tropical fruits from Brazil,” Food Chemistry, vol. 121, no. 4, pp. 996–1002, 2010. View at: Publisher Site | Google Scholar
  10. M. M. Root, M. C. Mcginn, D. C. Nieman et al., “Combined fruit and vegetable intake is correlated with improved inflammatory and oxidant status from a cross-sectional study in a community setting,” Nutrients, vol. 4, no. 1, pp. 29–41, 2012. View at: Publisher Site | Google Scholar
  11. J. Pérez-Jiménez, S. Arranz, M. Tabernero et al., “Updated methodology to determine antioxidant capacity in plant foods, oils and beverages: extraction, measurement and expression of results,” Food Research International, vol. 41, no. 3, pp. 274–285, 2008. View at: Publisher Site | Google Scholar
  12. R. Apak, M. Özyürek, K. Güçlü, and E. Çapanoğlu, “Antioxidant activity/capacity measurement.1. Classification, physicochemical principles, mechanisms, and electron transfer (ET)-based assays,” Journal of AOAC International, vol. 64, pp. 997–1027, 2016. View at: Publisher Site | Google Scholar
  13. L. P. Santos, D. R. Morais, N. E. Souza, S. M. Cottica, M. Boroski, and J. V. Visentainer, “Phenolic compounds and fatty acids in different parts of Vitis labrusca and V. vinifera grapes,” Food Research International, vol. 44, no. 5, pp. 1414–1418, 2011. View at: Publisher Site | Google Scholar
  14. W. Brand-Williams, M. E. Cuvelier, and C. Berset, “Use of a free radical method to evaluate antioxidant activity,” LWT—Food Science and Technology, vol. 28, no. 1, pp. 25–30, 1995. View at: Publisher Site | Google Scholar
  15. I. F. F. Benzie and J. J. Strain, “The ferric reducing ability of plasma (FRAP) as a measure of ‘antioxidant power’: the FRAP assay,” Analytical Biochemistry, vol. 239, no. 1, pp. 70–76, 1996. View at: Publisher Site | Google Scholar
  16. F. Shahidi and M. Naczk, Food Phenolics: Sources, Chemistry, Effects and Applications, Technomic Publishing Company, Lancaster, UK, 1995.
  17. R. G. Woisky and A. Salatino, “Analysis of propolis: Some parameters and procedures for chemical quality control,” Journal of Apicultural Research, vol. 37, no. 2, pp. 99–105, 1998. View at: Publisher Site | Google Scholar
  18. D. H. Lees and F. J. Francis, “Standardization of pigment analyses in cranberries,” HortScience, vol. 7, pp. 83–84, 1972. View at: Google Scholar
  19. E. G. Bligh and W. J. Dyer, “A rapid method of total lipid extraction and purification,” Canadian Journal of Physiology and Pharmacology, vol. 37, no. 8, pp. 911–917, 1959. View at: Google Scholar
  20. L. Hartman and R. C. A. Lago, “Rapid preparation of fatty acid methyl esters from lipids,” Laboratory Practice, vol. 22, no. 7, pp. 475–476, 1973. View at: Google Scholar
  21. J. D. Joseph and R. G. Ackman, “Capillary column gas chromatography method for analysis of encapsulated fish oil and fish oil ethyl esters: collaborative study,” Journal of AOAC International, vol. 75, pp. 488–506, 1992. View at: Google Scholar
  22. J. V. Visentainer, “Aspectos analíticos da resposta do detector de ionização em chama para ésteres de ácidos graxos em biodiesel e alimentos,” Química Nova, vol. 35, no. 2, pp. 274–279, 2012. View at: Publisher Site | Google Scholar
  23. F. Fratianni, M. Tucci, M. D. Palma, R. Pepe, and F. Nazzaro, “Polyphenolic composition in different parts of some cultivars of globe artichoke (Cynara cardunculus L. var. scolymus (L.) Fiori),” Food Chemistry, vol. 104, no. 3, pp. 1282–1286, 2007. View at: Publisher Site | Google Scholar
  24. A. C. Volp, I. R. T. Renhe, K. B. Barra, and P. C. Stringueta, “Flavonóides antocianinas: características e propriedades na nutrição e saúde,” Revista brasileira de nutrição clínica, vol. 23, pp. 141–149, 2008. View at: Google Scholar
  25. A. F. Faria, M. C. Marques, and A. Z. Mercadante, “Identification of bioactive compounds from jambolão (Syzygium cumini) and antioxidant capacity evaluation in different pH conditions,” Food Chemistry, vol. 126, no. 4, pp. 1571–1578, 2011. View at: Publisher Site | Google Scholar
  26. P. S. Benherlal and C. Arumughan, “Chemical composition and in vitro antioxidant studies on Syzygium cumini fruit,” Journal of the Science of Food and Agriculture, vol. 87, no. 14, pp. 2560–2569, 2007. View at: Publisher Site | Google Scholar
  27. C. Vasco, J. Ruales, and A. Kamal-Eldin, “Total phenolic compounds and antioxidant capacities of major fruits from Ecuador,” Food Chemistry, vol. 111, no. 4, pp. 816–823, 2008. View at: Publisher Site | Google Scholar
  28. D. Donno, R. Boggia, P. Zunin et al., “Phytochemical fingerprint and chemometrics for natural food preparation pattern recognition: an innovative technique in food supplement quality control,” Journal of Food Science and Technology, vol. 53, no. 2, pp. 1071–1083, 2016. View at: Publisher Site | Google Scholar
  29. T. Suzuki, S. Someya, F. Hu, and M. Tanokura, “Comparative study of catechin compositions in five Japanese persimmons (Diospyros kaki),” Food Chemistry, vol. 93, no. 1, pp. 149–152, 2005. View at: Publisher Site | Google Scholar
  30. J. M. Monteiro, U. P. De Albuquerque, E. L. De Araújo, and E. L. C. De Amorim, “Taninos: uma abordagem da química à ecologia,” Química Nova, vol. 28, no. 5, pp. 892–896, 2005. View at: Publisher Site | Google Scholar
  31. N. J. Kang, K. W. Lee, D. E. Lee et al., “Cocoa procyanidins suppress transformation by inhibiting mitogen-activated protein kinase kinase,” Journal of Biological Chemistry, vol. 283, no. 30, pp. 20664–20673, 2008. View at: Publisher Site | Google Scholar
  32. I. Rodríguez-Ramiro, S. Ramos, L. Bravo, L. Goya, and M. Á. Martín, “Procyanidin B2 induces Nrf2 translocation and glutathione S-transferase P1 expression via ERKs and p38-MAPK pathways and protect human colonic cells against oxidative stress,” European Journal of Nutrition, vol. 51, no. 7, pp. 881–892, 2012. View at: Publisher Site | Google Scholar
  33. H. Wang, F. Feng, B.-Y. Zhuang, and Y. Sun, “Evaluation of hepatoprotective effect of Zhi-Zi-Da-Huang decoction and its two fractions against acute alcohol-induced liver injury in rats,” Journal of Ethnopharmacology, vol. 126, no. 2, pp. 273–279, 2009. View at: Publisher Site | Google Scholar
  34. C. Romero, M. V. Ruiz-Méndez, and M. Brenes, “Bioactive compounds in virgin olive oil of the pdo montoro-adamuz,” JAOCS, Journal of the American Oil Chemists' Society, pp. 665–672, 2016. View at: Publisher Site | Google Scholar
  35. D. Kromhout, S. Yasuda, J. M. Geleijnse, and H. Shimokawa, “Fish oil and omega-3 fatty acids in cardiovascular disease: do they really work?” European Heart Journal, vol. 33, no. 4, pp. 436–443, 2012. View at: Publisher Site | Google Scholar
  36. C. Moreno, A. Macías, A. Prieto, A. de la Cruz, T. González, and C. Valenzuela, “Effects of n-3 polyunsaturated fatty acids on cardiac ion channels,” Frontiers in Physiology, vol. 3, article 245, 2012. View at: Publisher Site | Google Scholar
  37. A. P. Simopoulos, “Omega-3 fatty acids in inflammation and autoimmune diseases,” Journal of the American College of Nutrition, vol. 21, no. 6, pp. 495–505, 2002. View at: Publisher Site | Google Scholar
  38. R. De Caterina, R. Madonna, R. Zucchi, and M. T. La Rovere, “Antiarrhythmic effects of omega-3 fatty acids: from epidemiology to bedside,” American Heart Journal, vol. 146, no. 3, pp. 420–430, 2003. View at: Publisher Site | Google Scholar
  39. DACH, Referenzwerte Für Die Nährstoffzufuhr, Umschau/Braus GmbH, Frankfurt, Germany, 1st edition, 2002.

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