Polyphenol and Ellagitannin Constituents of Jabuticaba (Myrciaria

Article pubs.acs.org/JAFC

Polyphenol and Ellagitannin Constituents of Jabuticaba (Myrciaria caulif lora) and Chemical Variability at Different Stages of Fruit Development Luciane Dias Pereira,† Joaõ Marcos Gonçalves Barbosa,† Antonio Jorge Ribeiro da Silva,‡ Pedro Henrique Ferri,† and Suzana Costa Santos*,† †

Instituto de Química, Universidade Federal de Goiás, 74690-900, Goiânia, Goiás Brazil Núcleo de Pesquisas de Produtos Naturais, Centro de Ciências da Saúde, Universidade Federal do Rio de Janeiro, 21941-590, Rio de Janeiro, Rio de Janeiro, Brazil

‡

S Supporting Information *

ABSTRACT: A new ellagitannin named cauliflorin (1), seven known hydrolyzable tannins (2−8), and six known phenolics (9− 14) were isolated from jabuticaba. Compounds 2−8 had not been previously isolated from M. caulif lora fruits. The jabuticaba fruit was analyzed at four developmental stages for ellagitannins 1, 3, 7, and 8, phenolic acids 11 and 12, anthocyanins, organic acids, and sugars via HPLC-UV-DAD and NMRq. The content of ellagitannins and organic acids declined during fruit development, whereas at full ripeness sugar and anthocyanin levels underwent a sharp increase and were mainly constituted by fructose and cyanidin-3-O-glucose, respectively. Ellagitannins’ profile varied considerably among fruit tissues, with pedunculagin (3), castalagin (7), and vescalagin (8) mostly concentrated in jabuticaba seeds, whereas cauliflorin (1) and anthocyanins accumulated in the peels. Changes in jabuticaba’s phenolic compound contents were mostly influenced by fruit part (peel, pulp, and seed) rather than by degree of ripeness. KEYWORDS: Myrciaria caulif lora, jaboticaba, ellagitannin, flavonoids, anthocyanins, structural elucidation, NMR, HPLC-DAD

■

INTRODUCTION The jabuticabeira [Myrciaria caulif lora (Mart.) O. Berg and M. jaboticaba (Vell.) O. Berg] is widely known as “Brazilian grape tree”, and its dark-colored, acid sweet fruit called jabuticaba or jaboticaba grows directly on its main trunks and stems. Decoction of the fruit’s dark-purple peels is used in folk medicine as treatment for hemoptysis, throat inflammation, cough, bronchitis, and asthma, whereas the bark and leaves are used to treat diarrhea and dysentery.1 The jabuticaba has become a new source of bioactive compounds, as shown by the increasing attention it has received in the past few years; 83 reports regarding its biological activities and chemical profile have been published from 2010 to 2016, according to SciFinder database statistics. Among the health benefits confirmed for jabuticaba are radical scavenger capacity, antiproliferative effects against tumor cell lines, anti-inflammatory, anticancer, antibacterial, and antidiabetic activities, as well as protection against hepatic damage and collagen degradation.1 Most biological activities are due to high polyphenol contents in both Myrciaria species, which include compounds such as flavonoids, anthocyanins, phenolic acids, and ellagitannins. Regarding jabuticaba’s phytochemical composition, few studies have focused on isolating and identifying phenolic compounds;2,3 instead, they tend to concentrate on suggesting the structure of compounds, mainly ellagitannins, based on LC/MS studies.4−6 Jabuticaba is consumed as fresh fruits, but due to rapid fruit decay which takes place in 2−3 days, producing jams, ice creams, juices, liqueurs, distillates, and wines is an alternative to prevent postharvest losses.1 Research on exotic fruit wines has © XXXX American Chemical Society

recently intensified as a form of assigning value to regional fruits and stimulating their production ;7 however, the quality of jabuticaba’s processed products is still of concern as undesirable chemical changes may occur during fruit processing and storage.1,8 Throughout the development and ripening processes, fruits also undergo biochemical, physiological, and structural changes that determine their final quality, possibly affecting their health properties and industrial use. Even though changes in chemical compounds during jabuticaba’s maturation have been reported previously, showing variations in carbohydrates, organic acids, and phenolic compounds,5,9,10 it is important to emphasize the lack of information concerning qualitative and quantitative aspects of individual ellagitannins during this fruit’s maturation. Therefore, the aim of this study was to isolate and identify ellagitannins and other phenolic compounds from seeds and peels as well as quantify isolated phenolics by HPLC-PDA in three tissues of jabuticaba fruits in four developmental stages. The quantitative 1H NMR technique was applied to measure sugars and organic acids amounts during jabuticaba’s development. To assess the influence of fruit parts and maturation degree on chemical variability, chemical constituents were submitted to canonical redundancy (RDA) and hierarchical cluster (HCA) analyses. These were carried out to detect sample distribution patterns and to identify which chemical Received: June 29, 2016 Revised: January 11, 2017 Accepted: January 31, 2017

A

DOI: 10.1021/acs.jafc.6b02929 J. Agric. Food Chem. XXXX, XXX, XXX−XXX

Article

Journal of Agricultural and Food Chemistry Table 1. NMR Data of Cauliflorin (acetone-d6)a δC position 1 2 3 4 5 6 6 tergalloyl A ring 1′ 2′ 3′ 4′ 5′ 6′ 7′ tergalloyl B ring 1′ 2′ 3′ 4′ 5′ 6′ 7′ tergalloyl C ring 1′ 2′ 3′ 4′ 5′ 6′ 7′ a

α-anomer 94.0 74.2 73.3 73.5 67.5 65.0

114.3 125.9 107.5 145.8 136.3 145.3b 168.2 121.3 133.1 112.1 148.1 142.6 145.4b 167.8 113.0 140.5 136.6 143.7 144.3 110.0 163.2

δH (J, Hz) β-anomer

α-anomer

99.0 77.1 75.5 73.8 72.0 65.0

5.19 3.57 3.89 4.81 4.42 5.05 3.76

d (3.7) dd (9.5, 3.7) t (9.5) t (9.5) ddd (9.5, 6.4, 1.0) dd (13.0, 6.4) d (13.0)

β-anomer 4.60 3.34 3.68 4.86 3.91 5.07 3.82

d (7.8) dd (9.3, 7.8) t (9.3) t (9.3) m dd (13.0, 6.4) dd (13.0)

6.73 s, 6.74 s

6.92 s, 6.93 s

6.91 s

From HSQC, HMBC, and COSY NMR experiments. bInterchangeable signals. October 2014. Fruits were sorted into four maturity stages according to external color: immature green (IG), breaker turning purple (BR), light purple semiripe (SR), and dark purple full-ripe (FR). Part of the berries was washed with running water and manually separated into its components (peel, pulp, and seeds). Berries and their separated parts were divided into samples that were evaluated as three replications of 100 g each. All samples were blended with 50 mL of distilled water and freeze dried. Dried samples were stored at −18 °C until analysis. Extraction and Isolation of Phenolic Compounds. Fresh seeds (1.8 kg) and peels (4.2 kg) were separately homogenized in a blender with water and then exhaustively extracted with 50% acetone using an overhead stirrer apparatus at room temperature. The acetone was removed under reduced pressure, and the suspended aqueous extract was filtered. Following, a liquid−liquid extraction with ethyl acetate (ETAC) was carried out. The combined organic phases were evaporated to yield 0.63 and 4.34 g of ETAC extracts from seed and peel, respectively. Aqueous layers were freeze dried and solubilized with methanol (MEOH), furnishing 21.45 and 56.56 g of MEOHsoluble fraction from seed and peel, respectively. The seed’s MEOHsoluble fraction was subjected to Diaion HP-20 column chromatography (27 × 4 cm i.d.) and eluted with a decreasing polarity gradient of H2O/MEOH. Following TLC analysis, eight main fractions were combined (SM1−SM8). SM4 (1.1 g) and SM7 (0.5 g) were separately applied to Sephadex LH-20 CC (27 × 1.8 cm i.d.), eluting with an increasing polarity gradient of CHCl3/ETOH followed by ETOH/ MEOH, to give compounds 1 (40 mg) from SM7 and 3 (321 mg), 7 (209 mg), and 8 (63 mg) from SM4. The seed’s ETAC extract was loaded onto a Sephadex LH-20 column (27 × 4 cm i.d.) and eluted with gradients of CHCl3/ETOH and ETOH/MEOH to give

constituents are able to distinguish between these sample groups. In addition, chemical variation partitioning was employed for different sources of assumed influence, i.e., fruit parts, ripeness, and genetic/environmental factors.

■

MATERIALS AND METHODS

Solvents, Materials, and Standards. HPLC gradient-grade acetonitrile was purchased from J.T. Baker (Mallinckrodt Baker Inc., Mexico). HPLC-grade H2O was prepared with a Milli-Q water system (Millipore Inc., Milli-Q, USA). Solvents for NMR were purchased from Cambridge Isotope Laboratory Inc. (Andover, MA). All other solvents and reagents used were of analytical grade. Column chromatography was run using Diaion HP-20 and Sephadex LH-20 (Sigma-Aldrich Chemical Co., St. Louis, MO, USA). Analytical TLC was carried out with Silica gel 60 F254 plates (Merck, Darmstadt, Germany) using formic acid−ethyl formiate−toluene (1:7:1) as mobile phase. TLC spots were visualized by spraying plates with a 1% ethanolic solution of ferric chloride in HCl (0.1%) and UV light. Cyanidin 3-O-glucoside, gallic acid, ellagic acid, and TMSP-2,2,3,3-D4 (sodium-3-trimethylsilyl-propionate) were purchased from Sigma Chemicals Co. (St. Louis, MO, USA). Plant Material for Isolation of Phenolic Compounds. Cultivated M. caulif lora ripe fruits were collected at Jabuticabal Farm (S 16°49′53″, W 49°14′45″), Goiás State, Brazil, in October 2011. Approximately 8 kg of fresh fruits was washed with running water; seeds and peels were manually separated. Plant Material for Comparison of Developmental Stages. Cultivated M. cauliflora fruits were collected at Jabuticabal Farm in B


Article

Journal of Agricultural and Food Chemistry compounds 2 (73 mg), 4 (36 mg), and 11 (31 mg). The peel’s MEOH-soluble fraction was separated into two 28 g portions. These were submitted to column chromatography via Diaion HP-20 (27 × 4 cm i.d.) with MEOH/H2O step-gradient elution (0−100%) to yield five fractions (PM1−PM5). Fraction PM2 (8.0 g) was dissolved in water (100 mL) and extracted with ETAC (60 mL × 15); after solvent evaporation, this fraction (0.6 g) was subjected to Sephadex LH-20 CC (27 × 2 cm i.d.) and eluted with a stepwise gradient of CHCl3/ ETOH (7:3 to 0:10) followed by ETOH/MEOH (9:1 to 1:9) to afford compounds 5 (9 mg), 12 (8 mg), and 14 (14 mg). The peel’s ETAC extract was chromatographed on a 28 × 4 cm i.d. Sephadex LH20 column eluting with the same gradient of CHCl3/ETOH followed by ETOH/MEOH to give compounds 6 (46 mg) and 9 (124 mg) and a mixture of 10, 11, and 13 (319 mg). Cauliflorin (1). 1 is a white amorphous powder; UV (in mobile phase at tR 18.2 and 21.5 min) λmax (nm) 214, 278; ESI-TOF MS m/z 631.0585 [M − H]− (calcd for C27H19O18, 631.0577). 1H and 13C NMR data are given in Table 1. Preparation of Extracts for Comparison of Developmental Stages. For HPLC analysis, freeze-dried samples of whole fruits and parts (0.5 g) were homogenized with 5.0 mL of MEOH:H2O (8:2) in a test tube and sonicated for 15 min. The extract was separated from the solid residue by centrifuging at 2000g for 5 min and transferred to a 10.0 mL volumetric flask. The same procedure was repeated twice with 3.0 and 2.0 mL of MEOH 80% for 15 min each. Extracts were combined in a final 10.0 mL volume and prepared in triplicate. For 1H NMR quantification, freeze-dried samples of whole fruits (0.2 g) were homogenized with 10.0 mL of Milli-Q water in a test tube and sonicated for 30 min at 50 °C. The extract was separated from the solid residue by centrifuging at 2000g for 15 min and transferred to a 25.0 mL volumetric flask. The same procedure was repeated twice with 10.0 and 5.0 mL of Milli-Q water for 15 min each. Extracts were combined in a final 25.0 mL volume and freeze dried. HPLC Analysis. Polyphenols were quantified by a Shimadzu LC10AVP system (Shimadzu Corp., Japan) equipped with two LC10ADvp solvent delivery units connected to a photodiode array UV− vis detector (SPD-10AVvp) and a LiChrospher 100 RP-18 (5 μm) column, 25 cm × 0.4 cm i.d. (Merck Millipore, Billerica, MA, USA). All sample extracts were filtered through a 0.2 μm PTFE filter prior to analysis. The binary mobile phase consisted of acetonitrile (solvent A) and 0.01 M H3PO4: 0.01 M KH2PO4 (solvent B). The elution profile was as follows: 0−15 min, 7−10% A in B; 15−20 min, 10−12% A in B; 20−25 min, 12−14% A in B; held isocratic for 10 min; 35−40 min, 14−18% A in B; maintained for 3 min; then increased to 25% A over 7 min; changed to initial conditions in 5 min followed by a 5 min reequilibration. Analyses were conducted using a 1.0 mL/min flow rate and 20 μL sample injection volume. Simultaneous monitoring was performed at 216 nm to quantify ellagitannins and gallic acid and 370 nm for anthocyanins and ellagic acid. UV spectra were recorded between 190 and 370 nm. Compounds in extracts were identified by comparing their RT values and UV spectra with those of commercial standards (cyanidin3-O-glucoside, gallic acid, and ellagic acid) and isolated compounds (cauliflorin (1), pedunculagin (3), castalagin (7), and vescalagin (8)). The calibration curve of cyanidin-3-O-glucoside was used to quantify delphinidin-3-O-glucoside. Quantification was based on the measured integration area applying the calibration equation of the corresponding standard, and values were expressed as milligrams in 10 g of dry fruit or fruit part (peel, pulp, and seed). Spectroscopy Measurements. ESI-TOF MS spectra were recorded on a Bruker microTOF instrument by direct infusion in negative ion mode. All NMR experiments were recorded on a Bruker Avance III 500 spectrometer operating at 500.13 MHz for 1H and at 125 MHz for 13C. For structural elucidation, compounds were dissolved in acetone-d6, acetone-d6 + D2O, or methanol-d4 using TMS as chemical shift reference (δ = 0 ppm). Conventional pulse sequences were applied to acquire 1H and 13C spectra as well as 2D NMR experiments such as DQF-COSY, 1H−13C HSQC, and 1H−13C HMBC spectra. For quantification of sugars (sucrose, glucose, and fructose) and organic acids (citric acid and malic acid), freeze-dried

extracts were dissolved in 2.0 mL of HCl solution (pH 1.0), centrifuged for 15 min, and filtered through a 2.0 μm PTFE filter. The dissolved extract (0.5 mL) was placed in a 5 mm NMR tube, and 0.1 mL of TMSP solution (10.5 g/L of TMSP and 70% v/v D2O) was added as internal standard for quantitative analysis and internal chemical shift reference (δ = 0 ppm). The following parameters were applied to quantitative 1H NMR spectra: the spectral window was 10 ppm, and data was collected into 65 k data points after 48 scans; the recycle delay was 5 s and had a flip angle of 90°, with an acquisition time of 4.06 s at a fixed temperature of 25 °C. A NOESYGPPR 1D pulse sequence (Bruker library) was applied to suppress the residual water signal by selective low-power irradiation at the water frequency during the relaxation delay; this pulse sequence showed no effect on the area of other signals. Data was analyzed by the TopSpin 2.1 software (Bruker BioSpin Corp., MA, USA). Five metabolites were quantified by measuring the peak area ratio of their signals in the 1H NMR spectrum relative to TMSP. Chemometric Data Analysis. Multivariate statistical analysis was performed using CANOCO (Canonical Community Ordination) version 5 (Biometrics, The Netherlands, 2012). Chemical constituents were ordered in a response data matrix with rows = samples and columns = chemical variables. Fruit factors were ordered in an explanatory data matrix with rows = samples and columns = degree of fruit maturity (IG, BR, SR, FR) and fruit part (peel, pulp, and seed). Preliminary analyses applied the default options of the detrended canonical analysis (DCA) to CANOCO to assess the magnitude of dispersion in the response matrix in relation to the explanatory matrix along the first ordination axis (i.e., gradient length in standard deviation units, SD). In this study, DCA estimated the compositional gradient in the response data to be shorter than 2.0 SD units; thus, canonical redundancy analysis (RDA) was the appropriate ordination method to perform a linear direct gradient analysis.11 An unrestricted Monte Carlo permutation test (9999 permutations) was used to evaluate the significance of canonical eigenvalues. In all RDAs, the variance inflation factor (VIF) with sequential Bonferroni technique was used to guide the selection of explanatory variables, avoiding multicollinearity in multivariate regressions. VIF values > 20 are considered strongly multicollinear.11 The hierarchical cluster analysis (HCA) was used to detect natural groupings from RDAs and their intra- and intergroup relations. All hierarchical clusterings used Euclidean distance with variance minimization by Ward’s technique.12 In addition to the above-mentioned techniques, partitioning of the total variance response matrix was obtained by partial RDAs (pRDAs) using the explanatory matrix reordered in two or three groups of variables.13,14 The significant terms in each group were selected by a progressive variable selection procedure available in CANOCO with the significance level set by the Bonferroni correction and using the VIF variable as criterion to assess multicollinearity in multiple regressions and to reduce error type I.11,15 Partitioning of total data variance of the response matrix resulted in different fractions of data variation with or without overlap between the different groups of explanatory variables. Prior to the multivariate analysis, the data was preprocessed: a response matrix was transformed to log(x + 1), centered, and autoscaled in most analyses, whereas the explanatory matrix was centered and autoscaled. Average multiple comparisons were established by ANOVA using SAS GLM analyses (Statistical Analysis System, SAS Institute Inc., Cary, NC, 1996). All data was checked for homoscedasticity with Hartley’s test. Whenever heteroscedasticity was observed, the variable was angular or rank transformed. Whenever a difference was established in ANOVA, a posthoc Tukey test was performed. p Values below 0.05 were regarded as significant.

■

RESULTS AND DISCUSSION Structure Elucidation. Cauliflorin (1) has the molecular formula of C27H20O18 on the basis of its ESI-TOF MS at m/z 631.0585 [M − H]−. The 1H and 13C NMR spectra (Figures S1 C


Article

Journal of Agricultural and Food Chemistry

Figure 1. Structures of compounds 1−14 isolated from Myrciaria cauliflora fruits.

between δH 6.91 and δH 6.93 and their respective carbons at δC 110.0 and δC 112.1, which are not typical of HHDP groups.20 The evaluation of the 1H−13C long-range correlation map from HMBC NMR experiment (Figure S5 and Table S1) revealed correlations between singlets δH 6.92 and δH 6.93 with carbons δC 121.3, 142.6, and 148.1 and carbonyl δC 167.8; the latter also correlates with the hydrogens attached to the C-6 of α and β glucoses. The unusual higher chemical shifts for C-1′, C-4′, and C-5′ of the esterified phenoyl ring (B) linked to O-6 of the glucose core (Table 1), in comparison to common HHDP groups, suggest that the third aromatic ring (C) was attached to one of the oxygens of this unit. The aromatic hydrogens at δH 6.91 exhibited 1H−13C long-range HMBC correlations to carbons at δC 113.0, 140.5, 143.7, 144.3, and 163.2. The low chemical shift of carbonyl indicates a lactone function linked to the third phenoyl ring (C), which is in agreement with compounds alnusiin (2) and praecoxin C, in which chemical shifts for lactone carbonyl groups occurred at δC 163 ppm.20 The direction of the lactone ring was determined by comparing the 13C NMR signals of 1 with those of praecoxin C and alnusiin A.20,21 The comparison of carbon signals from the B ring of the depsidone-forming valoneoyl group (C-4′, C5′, and C-6′: δC 151.7, 135.5, and 148.5 in praecoxin C20) and of the depsidone-forming tergalloyl group (C-4′, C-5′, and C6′: δC 148.2, 142.8, and 145.2 in alnusiin A21) with compound 1 (Table 1) showed consistent similarity of B-ring carbon shifts of 1 with a depsidic tergalloyl moiety; therefore, the depsidone linkage was assigned at C-6′ and the ether linkage at C-5′ of the B ring. Consequently, 1 was characterized as 4,6-O-tergalloyl-Dglucose and named cauliflorin (Figure 1). To our knowledge,

and S2) showed a mixture of α and β anomers due to the absence of an ester group at the anomeric center. This was evident by the presence in the 13C NMR spectrum of two signals for anomeric carbons at δC 94.0 (C-1α) and δC 99.0 (C1β) as well as of two visible doublets at δH 5.19 (J = 3.7 Hz) and δH 4.60 (J = 7.8 Hz) in the 1H NMR spectrum attributed to α and β anomeric hydrogens, respectively. The fact that the signals of anomeric hydrogens are more shielded, compared with those of pedunculagin (3) and alnusiin (2), as well as the large difference in the chemical shifts between anomeric carbons (Δδβ−α = 5.0) already indicate that the hydroxyl in position 2 is also not esterified.16,17 All other glucose signals were present in pairs in both NMR spectra. Glucose hydrogen signals of both anomers were identified by the 1H−1H correlation COSY NMR experiment (Figure S3), and coupling constants indicated the 4C 1 conformation of the glucopyranose ring.18 Hydrogens H-2 and H-3 from both anomers were shielded and resonated between δH 3.34 and 3.89, demonstrating no esterification at these positions, in contrast to deshielded hydrogens H-4 and H-6, thus indicating acyl groups at O-4 and O-6. The two H-6 hydrogens in each anomer exhibited a large difference between their chemical shifts (ΔδH 1.25 and 1.29), which is typical of ellagitannins with an HHDP (or the HHDP part of a valoneoyl or tergalloyl) group located at O-4 and O-6.19 The aromatic hydrogen region of the 1H NMR spectrum showed four one-hydrogen singlets at δH 6.73, 6.74, 6.92, and 6.93 and one two-hydrogen singlet at δH 6.91, indicating three aromatic hydrogens for each anomer, which points to the existence of an aromatic ring attached to the HHDP group. This is confirmed by the deshielded signals of hydrogens D


Article


Figure 2. 1H NMR spectrum of jabuticaba’s fruit extracts: A, IG stage (immature green); B, FR stage (full ripe) (500 MHz, HCl/H2O/D2O solution, pH 1.0).

the present study is the first report of this ellagitannin in the literature. The other isolates were alnusiin (2), pedunculagin (3), strictinin (4), casuarictin (5), 1,2,3,4,6-penta-O-galloyl-β-Dglucose (6), castalagin (7), vescalagin (8), quercitrin (9), myricitrin (10), gallic acid (11), protocatechuic acid (12), ellagic acid (13), and depside 2-O-(3,4-dihydroxybenzoyl)2,4,6-trihydroxyphenylacetic acid (14) (Figure 1), whose spectral data agreed with reported values. Spectral data of the known isolated ellagitannins are described in the Supporting Information. The phytochemical study of jabuticaba’s seeds and peels allowed the isolation and structural elucidation of 15

compounds by NMR spectroscopy. Eight substances were isolated and identified for the first time for M. caulif lora: a new ellagitannin 1 along with seven known hydrolyzable tannins 2− 8. In a previous study, which applied liquid chromatography coupled with a mass detector (LC-TOF-MS), the existence of ellagitannins 3−5 was suggested for fruits of this species.4 In the same report, four other ellagitannins were also suggested, but they were not identified in the present study. Another ellagitannin profile, obtained by LC-MS, has been suggested elsewhere,5 but it was not consistent with the present study or with reports by Wu and collaborators.3,4 Rare ellagitannin alnusiin (2), also named alnusnin B/alunusnin B,22 which contains in its structure the unusual component unit, tergalloyl E


Article


Figure 3. HPLC-PDA of jabuticaba fruit extracts: IG stage (A, 216 nm; B, 370 nm), FR stage (C, 216 nm; D, 370 nm). Compounds: 1, gallic acid; 2, vescalagin; 3, pedunculagin; 4, castalagin; 5, pedunculagin; 6, cauliflorin; 7, ellagic acid; 8, delphinidin-3-O-glucose; 9, cianydin-3-O-glucose.

ester group, has been isolated only from fruits of Alnus sieboldiana, Betulaceae,22,23 and Rosa roxburghii, Rosaceae.24 In biological tests, alnusiin demonstrated high antitumor activity against sarcoma-180 in vivo and inhibited lipid peroxidation at 88%, induced by NADPH and ADP in rat liver microsomes.25,26 Alnusiin affected lipid metabolism induced by hormones adrenaline, adrenocortical, and insulin in fat cells isolated from rat adipose tissue.27 This ellagitannin also showed an antihepatotoxic effect in lesions induced by CCl4.28 Changes in Organic Acids, Sugars, and Phenolic Compounds during Fruit Development. Jabuticaba fruits collected in four ripening stagesimmature green (IG), breaker turning purple (BR), light purple semiripe (SR), and dark purple full ripe (FR)were analyzed for organic acids (citric and malic acid) and sugars (sucrose, glucose, and fructose) via a quantitative NMR technique. Chemical shifts of selected signals and 1H NMR spectra are shown in Table S2 and Figure 2. Reversed-phase HPLC coupled with PDA detection was used to quantify phenolic compounds 1, 3, 7, 8, 11, 13, cyanidin-3-O-glucosside, and delphinidin-3-O-glucoside in fruit extracts. Retention times, UV data, and chromatograms are shown in Table S3 and Figure 3. Results obtained in the measurements of organic acids, sugars, and phenolic compounds in fruit extracts were subjected to multivariate statistical analysis of canonical redundancy (RDA). In the first exploratory analysis (Figure S7), which considered the four maturation stages, samples were ordered in three distinct groups, and fruits of the second and third stages (BR and SR) formed a single group. To confirm this trend, hierarchical cluster analysis (HCA) was used to detect natural groupings (Figure S8). HCA confirmed the existence of only three sample groups; there are no significant chemical differences between BR and SR stages that justify separating samples into four groups. A new RDA was performed considering as response matrix 12 samples × 13 chemical compounds and as explanatory matrix 12 samples × 1 factor with 3 categories (IG, BR + SR, FR). Results of ordination analysis via RDA (Figure 4) showed that the correlations between sets of response and maturation data (explanatory matrix) were higher in the first two canonical axes (0.9851 and 0.9213), and VIFs were considered low (VIF < 1.5), hence suggesting the absence of multicollinearity between variables.11 Monte Carlo permutation tests (999 permutations) showed highly significant results for the first two canonical axes (RDA1: F-Fischer = 65.8, p < 0.001; RDA2: F = 6.8, p < 0.004),

Figure 4. RDA ordination of the first two axes showing the distribution of jabuticaba sampling maturity stages (IG, BR, SR, FR). Chemical constituents of fruits are represented by long arrows from the origin. Red triangles represent cluster centroids. Values in brackets refer to the explained variance on each canonical axis.

signaling that variation patterns in the original arrays do not arise by chance. The sum of canonical eigenvalues was also highly significant (matrix trace response = 0.946; F = 61.0, p < 0.001).11,14,15 These results suggest a strong and significant association between the fruit’s chemical constituents and the three maturity stages (explanatory factor). According to Figure 4, the RDA1 axis shows the correlation between the degree of maturation and fruits’ chemical constituents. A reduction in RDA1 is associated with the green stage (IG), whose samples have higher levels of all acids (citric, malic, gallic, and ellagic), as well as the four ellagitannins, whereas an increase in RDA1 is correlated with the last stage (FR), during which sugar and anthocyanin contents are higher. A reduction in RDA2 values correlates with the intermediate maturation stage (BR + SR), especially with variables 3, 7, 8, and cyanidin-3-O-glucoside. Sugars and organic acids contribute appreciably to fruit flavor. Jabuticaba’s fraction of soluble carbohydrates consisted mainly of fructose, glucose, and sucrose (Table 2). Higher fructose levels throughout fruit development had already been reported by two previous studies on jabuticaba.5,29 Glucose and fructose levels increased mainly from the intermediate- to the full-ripe stage, 3- and 2.5-fold, respectively (Table 2), whereas sucrose concentration increased about 20 times from the IG stage until the end of maturation (FR), in agreement with a previous report.5 F


Article


transported in the phloem to fruits, where it can be accumulated and/or hydrolyzed into glucose and fructose by invertase, but glucose may also be formed by starch degradation which occurs in jabuticaba pulp cells during maturation.31 Therefore, during fruit ripening, the amount of glucose increases at a faster rate than fructose, so the ratio of glucose/fructose (G/F) will be proportional to the degree of maturity.32 In the maturation process of grapes from cultivar Autumn Seedless, there was a variation in the G/F ratio from 0.98 to 1.05.33 Similarly, jabuticaba’s G/F ratio also increased from 0.54 to 0.88 during fruit ripeness. The main nonaromatic organic acids analyzed in jabuticaba fruits were citric and malic acid (Table 2). These acids are responsible for the fruit’s total acidity and the final quality of jabuticaba wine. Unlike other studies,29,34 succinic acid was not detected in this work. This difference may be due to factors such as soil fertility, irrigation, solar intensity, cultivar, and temperature, which directly influence the content of organic acids in fruits.35 Throughout jabuticaba’s development, acid concentrations significantly decreased, and at the FR stage citric acid and malic acid levels were reduced by 48.7% and 76.4% of their initial content, respectively. Malate is used mostly as a source of energy for the fruit during maturation, so its levels decreased faster than those of the citrate.32 Cyanidin-3-O-glucoside and delphinidin-3-O-glucoside, anthocyanins responsible for peel color, were detected starting from the second growth stage. Both reached their maximum concentration at the end of ripening, when the fruit became dark purple (Table 2). These levels were similar to those obtained from jabuticaba grown in Brazil,5 12.3 and 2.35 mg/

Table 2. Contents of Sugars, Organic Acids (g/100g DW), Phenolic Acids, Ellagitannins, and Anthocyanins (mg/10g DW) in the Clustered Jabuticaba Fruitsa clusters variables sucroseb glucoseb fructosec citric acid malic acid gallic acid (11) vescalagin (8) castalagin (7) pedunculagin (3) cauliflorin (1)b delphinidin3-O-glucosideb,d cyanidin3-O-glucoside ellagic acid (13)

IG 1.05 14.97 27.57 27.75 4.36 2.18 37.27 33.91 54.26 27.66 0.00

± ± ± ± ± ± ± ± ± ± ±

0.23 1.18 0.20 1.23 0.15 0.37 6.10 3.14 6.10 8.42 0.00

BR + SR c c b a a a a a a a c

4.82 18.31 25.73 18.63 1.67 0.97 27.30 26.85 23.41 7.88 0.62

± ± ± ± ± ± ± ± ± ± ±

0.69 1.32 3.22 1.36 0.24 0.13 8.01 3.97 7.10 1.14 0.21

FR b b b b b b a a b b b

22.67 53.62 61.16 14.20 1.03 0.58 8.48 15.50 4.32 3.15 4.19

± ± ± ± ± ± ± ± ± ± ±

2.67 4.85 4.63 0.85 0.15 0.12 3.78 3.57 2.29 0.91 1.96

a a a c c b b b c c a

0.00 ± 0.00 c

3.54 ± 1.70 b

10.79 ± 0.55 a

4.35 ± 0.36 a

1.85 ±0 .17 b

1.54 ± 0.32 b

a

Average based on original data in triplicate. bRank transformed in ANOVA analysis. cAngular transformed in ANOVA analysis. d Calibration curve used: cyanidin-3-O-glucoside (after correction). Averages followed by the same letter in the rows did not share significant differences at 5% probability by Tukey’s test. IG, immature green; BR, breaker turning purple; SR, semiripe; FR, full-ripe stage.

Variations in sugar contents are influenced by the activities of enzymes related to the metabolism of carbohydrates, such as invertase and sucrose phosphate synthase.30 Sucrose is

Table 3. Average Concentrationsa (mg/10g DW) of Polyphenols in Jabuticaba Parts at Different Developmental Stages developmental stage compounds gallic acid (11)b

vescalagin (8)b

castalagin (7)b

pedunculagin (3)b

cauliflorin (1)b

delphinidin-3-O-glucosideb,c

cyanidin-3-O-glucoside

ellagic acid (13)b

fruit part peel seed pulp peel seed pulp peel seed pulp peel seed pulp peel seed pulp peel seed pulp peel seed pulp peel seed pulp

IG

BR

SR

FR

1.86 4.19 0.43 1.95 281.67 1.87

± ± ± ± ± ±

0.36 0.15 0.02 0.52 1.21 0.23

Ba Aa Cab Ba Aa Bc

0.93 2.53 0.52 0.90 134.00 10.30

± ± ± ± ± ±

0.02 0.49 0.03 0.26 1.01 0.43

Bab Ab Ca Cb Abc Ba

0.79 2.40 0.41 0.49 145.67 7.14

± ± ± ± ± ±

0.08 0.34 0.02 0.02 8.35 0.71

Bb Ab Cbc Cbc Aab Ba

0.49 2.36 0.35 0.41 95.27 2.46

± ± ± ± ± ±

0.07 Bc 0.22 Ab 0.01 Cc 0.03 Cc 10.90 Ac 0.10 Bb

255.33 5.22 33.50 45.18 7.53 12.58

± ± ± ± ± ±

3.22 1.03 9.07 0.41 0.94 2.06

Aa Bc Ba Aa Ca Aa

201.33 15.07 7.48 20.78 6.25 5.18

± ± ± ± ± ±

4.04 0.61 1.54 1.34 0.26 0.46

Ab Ba Bb Ab Cb Ab

214.67 12.14 2.06 13.33 6.11 1.50

± ± ± ± ± ±

13.65 Ab 0.88 Bab 0.15 Cc 0.65 Abc 0.49 Bb 0.09 Bc

175.67 7.70 2.23 11.75 2.39 2.12

± ± ± ± ± ±

16.92 Ac 0.18 Bb 0.27 Bc 2.50 Ac 0.13 Bb 0.53 Ac

3.00 ± 0.64 Bb

6.70 ± 0.44 0.85 ± 0.02

3.96 ± 0.22 Ba

3.13 ± 0.33 Ab 1.42 ± 0.09 b

1.64 ± 0.02 Bc 6.16 ± 0.81 a

3.92 ± 0.35 b

6.93 ± 0.86 b

27.2 ± 2.54 a

2.00 ± 0.09 Ba 5.59 ± 0.23 Ab 1.00 ± 0.04 Ca

1.73 ± 0.21 Ba 6.53 ± 0.40 Aa 0.90 ± 0.04 Cb

1.84 ± 0.15 Ba 5.82 ± 0.42 Aab 0.83 ± 0.03 Cb

a

Average based on original data in triplicate. bRank transformed in ANOVA analysis. cCalibration curve used: cyanidin-3-O-glucoside (after correction). Averages followed by the same capital letter in the columns and by the same lowercase letter in the rows did not share significant differences at 5% probability by the Tukey test. IG, immature green; BR, breaker turning purple; SR, semiripe; FR, full-ripe stage. G


Article


3). In green fruits, cauliflorin (1) levels were four times higher in the peel. In the breaker stage, its concentration decreased in the peel but showed a slight increase in the pulp, after which 1 reached minimum values on both parts at the end of ripening. Gallic acid (11), ellagic acid (13), pedunculagin (3), and vescalagin (8) were detected in all fruit parts; however, they occur in greater amounts in the seeds throughout fruit development. Reduction in their levels was the most common trend for these compounds, except for 7 and 8 contents in the pulp, which significantly increased in about 3- and 5-fold between the first and the second growth phase (Table 3). This variation may be due to the translocation of these substances between seed and pulp or to the biotransformation of pedunculagin (3) into 7 and 8.40 Castalagin (7) levels were higher than vescalagin (8) levels in seeds and pulp during most ripening stages, except in green seeds (Table 3). Castalagin (7) and vescalagin (8) are epimers whose single structural difference is the position of the hydroxyl at C-1; despite the structural similarity, these ellagitannins differ in chemical reactivity. Vescalagin (8) is more likely to undergo nucleophilic substitution reactions at C-1, and it is believed to be the precursor of several C-glycoside oligomers; it also conjugates with (+)-catechin, forming compounds known as flavano-ellagitannins.40 Therefore, in plant species with both compounds, castalagin (7) concentration is usually greater than vescalagin’s (8), as observed in the seeds of Myrciaria dubia, camu-camu, where the ratio of castalagin to vescalagin was approximately 6.0:4.0.47 As for the multivariate treatment, the data in Table 3 were ordered in two matrices: response matrix (36 samples × 8 chemical constituents) and explanatory matrix (36 samples × 2 factors: fruit part (peel, pulp, and seed) and degree of maturity (IG, BR, SR, and FR)). These two matrices were analyzed together via RDA to elucidate variation patterns of chemical constituents in samples regarding two factors: fruit part and degree of ripeness. Results of ordination analysis by RDA (Figure 5) indicated that correlations between chemical data (response matrix) and the two factors (explanatory matrix)

10g, but lower than those detected in fruit crops in the United States (43.3 and 81.0 mg/10g).36 Climatic factors such as temperature may be responsible for this difference. Anthocyanin accumulation is influenced by air temperature, particularly by day−night temperature variation.37 As has already been reported for strawberries, warm days and cool nights increase the production of pelargonidin and cyanidin.38 Anthocyanin levels in mature jabuticaba are crucial for wine production, since larger quantities of anthocyanins extracted from fruits will improve wine’s color properties and antioxidant effect.39 Ellagitannin (1, 3, 7, and 8) contents declined over fruit development and maturation stages, but pedunculagin (3) levels fell 57% from the first to the second growth phase, whereas castalagin (7) and vescalagin (8) showed no significant differences in the same period (Table 2). Until now, not all steps involved in the biosynthesis of C-glycoside ellagitannins were proved, but several experimental observations have subsidized the biosynthetic proposal with pedunculagin (3) as the key intermediate in the formation of these ellagitannins.40 Variations in pedunculagin (3) levels may, therefore, be due to its biotransformation in C-glycoside ellagitannins, which in turn can be combined to form dimers, trimers, and polymers. Pedunculagin (3) can also form dimers directly with Cglycoside ellagitannins, as reported in the leaves of English oak or Quercus robur.41 Furthermore, ellagitannin hydrolysis during the ripening process could be the cause of their reduction, but an increase in ellagic acid levels was not observed; on the contrary, a significant reduction of 58% in this acid occurred from the first to the second stage. Moreover, gallic acid also decreased by about 56%, which may be due to its use in the biosynthesis of C-glycoside ellagitannins. The decrease in tannin contents over the course of jabuticaba’s development is a common trend in other fruits such as myrtle and persimmon.42,43 It may result from the complexation of tannins with soluble pectins formed during fruit ripening,10 as well as from the initial dimerization of monomers and subsequent polymerization, generating high molecular weight tannins that can bind strongly to cell walls.41 Recent studies showed that a large proportion of ellagitannins remains bounded to cell walls and other macromolecules of the fruit.44 Therefore, the current trend for complete quantification of nonextractable ellagitannins involves acid hydrolysis of the freeze-dried fruit followed by HPLC-ESI-UV/MS/MS analysis of the released products.44 Changes of Phenolic Compounds during the Development of Jabuticaba Parts. The HPLC/PDA analysis was performed with compounds 1, 3, 7, 8, 11, 13, and anthocyanins to obtain the profile of phenolic compounds during fruit maturation in the different jabuticaba parts (peel, pulp, and seeds). Anthocyanins occur only in the jabuticaba peel, starting in the breaker toward the full-ripe stage (Table 3). The concentrations of both pigments increased by about four times until the final stage, but their levels were well below those obtained previously: 196.36 and 63.48 mg/10 g for cyanidin-3O-glucoside and delphinidin-3-O-glycoside, respectively.45 In the present study, phenolic compounds were not extracted by acidic solvents, since the use of organic or mineral acids could hydrolyze the ellagitannins. On the other hand, anthocyanins are less stable in the neutral medium,46 which could explain the lower levels obtained for these pigments. Even though cauliflorin (1) was isolated from the seeds, it was detected only in the crude extracts of pulp and peel (Table

Figure 5. RDA ordination of the first two axes showing the distribution of jabuticaba fruit parts samples (P, pulp; C, peel; S, seed) in four stages of maturity (IG, BR, SR, and FR). Chemical constituents of fruit parts are represented by long arrows from the origin. Degree of maturity is represented by red triangles, and blue triangles symbolize cluster centroids. Values in brackets refer to the explained variance on each canonical axis. H


Article

Journal of Agricultural and Food Chemistry were higher in the first two canonical axes (0.9920 and 0.7615). The VIFs were considered low (VIF < 1.5), suggesting there is no multicollinearity between variables.11 Monte Carlo permutation tests (999 permutations) showed highly significant results for the first two canonical axes (RDA1: F-Fischer = 134.0, p < 0.001; RDA2: F = 14.7, p < 0.001), signaling that variation patterns in the original arrays did not arise by chance. The sum of canonical eigenvalues was also highly significant (Trace response matrix = 0.946, F = 72.0, p < 0.001).11,14,15 These results suggest a strong and significant association between chemical constituents, on one hand, and fruit parts and ripening stages (explanatory factors), on the other. In Figure 5, the RDA1 axis showed a clear separation between seeds and the other two fruit parts (peel and pulp), whereas RDA2 described an ordering of samples according to the degree of fruit maturity. Thus, five sample classes were obtained, which were confirmed by the HCA through the scores of canonical axes from RDA (Figure S9). The difference between seeds and peel/pulp occurred mainly due to the massive concentration of castalagin (7) and vescalagin (8), as well as the high contents of gallic acid (11), ellagic acid (13), and pedunculagin (3) in seeds, similar to M. dubia.47 Separation between seeds of IG stage (class VI) from seeds of other stages (class V) is due to the high concentration of 3 in green seeds (Table 3). Peels were also separated into two groups: class III, which consists of peel samples at stages 2, 3, and 4, characterized by the presence of anthocyanins, whereas green peels were joined with pulps from stages 2, 3, and 4 in class I, which was related to cauliflorin (1). Green fruits pulps (class II) were also distinguished from pulps of other stages (class I) mainly by larger pedunculagin (3) levels and lower 7 and 8 contents (Table 3). Castalagin (7) and vescalagin (8) occur in high amounts in oak wood, Quercus robur and Q. petraea, and are found in grape wines aged in oak barrels. An analysis of Bordeaux red wine aged for 18 months in oak barrels showed these ellagitannins had a concentration of about 10 mg/L. These compounds contribute to wine’s various sensory properties, including taste and color.40 Glabasnia and Hofmann found that castalagin (7), vescalagin (8), and its derivatives impart an astringent mouthcoating sensation at remarkably low threshold concentrations in red wines aged in oak barrels.48 Vescalagin (8) can also influence wine color by reacting with anthocyanins and forming new pigments.49 Recently, a revision of the stereochemistry of the triphenoyl moiety of vescalagin and castalagin was conducted by DFT calculations of 1H and 13C NMR spectra, as well as TDDFT calculations of the ECD spectra of their desHHDP analogues. This study indicated that the triphenoyl moieties of these C-glycosidic ellagitannins exist in the (S,R) instead of (S,S) configuration.50 Results of the variation partitioning performed by partial redundancy analysis (pRDA) on the response matrix showed that the variation explained by fruit part ([a] in Table 4) was 70.8%, whereas only 13.4% of variation has been explained by degree of ripeness ([b]); a Venn diagram illustrating this variation partitioning is shown in the SI (Figure S10). These results suggest a stronger influence of the type of fruit tissue (peel, pulp, and seed) on the biosynthesis of phenolic compounds. It can be explained by the different functions of each fruit part: whereas anthocyanins color skins to promote a visual signal for the attraction of seed dispersal agents, tannins are involved in seed protection against pathogens and seed

Table 4. Summary of Variation Partitioning Using Partial RDA of Jabuticaba Fruit Parts Constituents at Different Developmental Stages effects and variables total effect fruit parts and maturity stage partial effects fruit parts fruit parts maturity stage maturity stage join effects fruit parts and maturity stage residuals

variation fraction

variation explained (%)a

F

Pb

[a + b + c]

84.2

32.0

0.001

maturity stage

[a]

70.8

67.2

0.001

fruit parts

[a + c] [b] [b + c]

70.8 13.4 13.4

40.1 8.5 1.7

0.001 0.001 0.131

covariables

[c]

0.0

[d]

15.8

Sum of canonical eigenvalues (l) divided by total inertia (1.0) × 100. b Probability based on Monte Carlo test (999 permutations). Predictors: Fruit parts = peel, pulp, and seed; developmental stages = IG, BR, SR, and FR. a

predators,51 similarly to achenes in strawberries and seeds of camu-camu, which accumulate defense-related phenolic compounds, including ellagitannins.47,52 Chemical variations in jabuticaba tissues were less explained by fruit development and ripening as the strong chemical shift was observed only between the first green stage (IG) and the later stages (BR, SR, FR) in each tissue (peel, pulp, and seed). The same distinction between green and ripe-stage metabolites (primary or secondary) had been reported in strawberry organs receptacle and achene; on the basis of metabolite difference, six stages were joined in two main groups for each organ.52 The unexplained variation in the data set reached 15.8% (see residuals, [d]), which indicates that chemovariations in phenolic compounds during fruit development and ripening may also be determined by other factors, such as climate (sunlight and temperature), water supply, variation in mineral nutrients, and hormone control.53 In conclusion, multivariate analysis of data obtained during fruit ripening ordered samples in three main groups; whereas the concentration of phenolic compounds and organic acids decreased, that of anthocyanins and sugars increased over the same period. The analysis of the separated fruit parts added a larger range of information, e.g., the distinction of seeds, which have higher levels of phenolic compounds, especially castalagin (7) and vescalagin (8). Additionally, the ordering of fruit parts in two major different groups exhibited marked chemical differences between the green stage of each tissue and later stages. This information opens up possibilities for new studies involving isolated tannins and anthocyanins and for the improvement of jabuticaba products.

■

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jafc.6b02929. NMR and mass spectra, NMR spectral data of ellagitannins and compounds in fruit extracts, HPLC I


Article


■

(14) Legendre, P.; Legendre, L. Numerical Ecology, 2nd ed.; Elsevier Science: Amsterdam, Netherlands, 2003; 853 pp. (15) Ter Braak, C. J. F.; Šmilauer, P. Canoco Reference Manual and User’s Guide: Software for Canonical Community Ordination (version 5.0); Microcomputer Power: New York, 2012; 496 pp. (16) Yoshida, T.; Hatano, T.; Okuda, T.; Memon, M. U.; Shingu, T.; Inoue, K. Spectral and chromatographic analyses of tannins. I. 13C Nuclear Magnetic Resonance spectra of hydrolysable tannins. Chem. Pharm. Bull. 1984, 32, 1790−1799. (17) Hatano, T.; Yoshida, T.; Shingu, T.; Okuda, T. 13C Nuclear Magnetic Resonance Spectra of Hydrolysable Tannins II. 1) Tannins forming anomer mixtures. Chem. Pharm. Bull. 1988, 36, 2925−2933. (18) Okuda, T.; Yoshida, T.; Hatano, T. New methods of analyzing tannins. J. Nat. Prod. 1989, 52, 1−31. (19) Yoshida, T.; Hatano, T.; Kuwajima, T.; Okuda, T. Oligomeric hydrolyzable tannins - their 1H NMR spectra partial degradation. Heterocycles 1992, 33, 463−482. (20) Hatano, T.; Yazaki, K.; Okonogi, A.; Okuda, T. Tannins of Stachyurus species. II. Praecoxins A, B, C and D, four new hydrolysable tannins from Stachyurus praecox leaves. Chem. Pharm. Bull. 1991, 39, 1689−1693. (21) Bao, L.-M.; Eerdunbayaer; Nozaki, A.; Takahashi, E.; Okamoto, K.; Ito, H.; Hatano, T. Hydrolysable tannins isolated from Syzygium aromaticum: structure of a new C-glucosidic ellagitannin and spectral features of tannins with a tergalloyl group. Heterocycles 2012, 85, 365− 381. (22) Tanaka, T.; Kirihara, S.; Nonaka, G.-I.; Nishioka, I. Tannins and Related Compounds. CXXIV. Five New Ellagitannins, Platycaryanins A, B, C, and D, and Platycariin, and a New Complex Tannin, Strobilanin, from the Fruits and Bark of Platycarya strobilacea SIEB et ZUCC., and Biomimetic Synthesis of C-Glycosidic Ellagitannins from Glucopyranose-Based Ellagitannins. Chem. Pharm. Bull. 1993, 41, 1708−1716. (23) Yoshida, T.; Memon, M. U.; Okuda, T. Alnusiin, a novel ellagitannin from Alnus sieboldiana fruits. Heterocycles 1981, 16, 1085− 1088. (24) Yoshida, T.; Chen, X.-M.; Hatano, T.; Fukushima, M.; Okuda, T. Tannins and related polyphenols of rosaceous medicinal plants. IV. Roxbins A and B from Rosa roxburghii fruits. Chem. Pharm. Bull. 1987, 35, 1817−1822. (25) Miyamoto, K.; Kishi, N.; Koshiura, R.; Yoshida, T.; Hatano, T.; Okuda, T. Relationship between sctructures and the antitumor activities of tannins. Chem. Pharm. Bull. 1987, 35, 814−822. (26) Okuda, T.; Kimura, Y.; Yoshida, T.; Hatano, T.; Okuda, H.; Arichi, S. Studies on the activities of tannins and related compounds from medicinal plants and drugs. I. Inhibitory effects on lipid peroxidation in mitochondria and microssomes of liver. Chem. Pharm. Bull. 1983, 31, 1625−1631. (27) Kimura, Y.; Okuda, H.; Okuda, T.; Yoshida, T.; Hatano, T.; Arichi, S. Studies on the activities of tannins and related compounds from medicinal plants and drugs. III. Effects of various tannins and related compounds on adrenocorticotropic hormon-induced lipolysis and insulin-induced lipogenesis from glucose in fat cells. Chem. Pharm. Bull. 1983, 31, 2501−2506. (28) Hikino, H.; Kiso, Y..; Hatano, T.; Yoshida, T.; Okuda, T. Antihepatotoxic actions of tannins. J. Ethnopharmacol. 1985, 14, 19− 29. (29) Lima, A. J. B.; Corrêa, A. D.; Dantas-Barros, A. M.; Nelson, D. L.; Amorim, A. C. L. Sugars, organic acids, minerals and lipids in jabuticaba. Rev. Bras. Frutic. 2011, 33, 540−550. (30) Srivastava, L. M. Fruit Development and Ripening. In Plant Growth and Development: Hormones and Environment; Academic Press: San Diego, CA, 2002; 413−429. (31) Barros, R. S.; Finger, F. L.; Magalhães, M. M. Changes in-nonstructural carbohydrates in developing fruit of Myrciaria jaboticaba. Sci. Hortic. 1996, 66, 209−215. (32) Sabir, A.; Kafkas, E.; Tangolar, S. Distribution of major sugars, acids and total phenols in juice of five grapevine (Vitis spp.) cultivars

data of phenolic compounds, biplot of RDA analysis, HCA dendograms, and Venn diagram (PDF)

AUTHOR INFORMATION

Corresponding Author

*Phone: + 55 62 3521 1008. E-mail: suzana.quimica.ufg@ hotmail.com. ORCID

Antonio Jorge Ribeiro da Silva: 0000-0002-7579-420X Suzana Costa Santos: 0000-0002-5583-3128 Funding

This work was supported by Conselho Nacional de ́ Desenvolvimento Cientifico e Tecnológico (CNPq, 470655/ 2012-7). L.D.P. received a scholarship from Fundaçaõ de Amparo à Pesquisa do Estado de Goiás (FAPEG). Notes

The authors declare no competing financial interest.

■

REFERENCES

(1) Wu, S.-B.; Long, C.; Kennelly, E. J. Phytochemistry and health benefits of jaboticaba, an emerging fruit crop from Brazil. Food Res. Int. 2013, 54, 148−159. (2) Reynertson, K. A.; Wallace, A. M.; Adachi, S.; Gil, R. R.; Yang, H.; Basile, M. J.; D’armiento, J.; Weinstein, I. B.; Kennelly, E. J. Bioactive Depsides and Anthocyanins from Jaboticaba (Myrciaria cauliflora). J. Nat. Prod. 2006, 69, 1228−1230. (3) Wu, S.-B.; Wu, J.; Yin, Z.; Zhang, J.; Long, C.; Kennelly, E. J.; Zheng, S. Bioactive and Marker Compounds from Two Edible DarkColored Myrciaria Fruits and the Synthesis of Jaboticabin. J. Agric. Food Chem. 2013, 61, 4035−4043. (4) Wu, S.-B.; Dastmalchi, K.; Long, C.; Kennelly, E. J. Metabolite profiling of jaboticaba (Myrciaria cauliflora) and other dark-colored fruit juices. J. Agric. Food Chem. 2012, 60, 7513−7525. (5) Alezandro, M. R.; Dubé, P.; Desjardins, Y.; Lajolo, F. M.; Genovese, M. I. Comparative study of chemical and phenolic compositions of two species of jaboticaba: Myrciaria jaboticaba (Vell.) Berg and Myrciaria caulif lora (Mart.) O. Berg. Food Res. Int. 2013, 54, 468−477. (6) Plaza, M.; Batista, A. G.; Cazarin, C. B. B.; Sandahl, M.; Turner, C.; Ö stman, E.; Maróstica, M. R., Jr. Characterization of antioxidant polyphenols from Myrciaria jaboticaba peel and their effects on glucose metabolism and antioxidant status: A pilot clinical study. Food Chem. 2016, 211, 185−197. (7) Duarte, W. F.; Dias, D. R.; Oliveira, J. M.; Teixeira, J. A.; Almeida e Silva, J. B.; Schwan, R. F. Characterization of different fruit wines made from cacao, cupuassu, gabiroba, jaboticaba and umbu. LWT Food Sci. Technol. 2010, 43, 1564−1572. (8) Fortes, G. A. C.; Naves, S. S.; Ferri, P. H.; Santos, S. C. Evaluation of chemical changes during Myrciaria caulif lora (Jabuticaba Fruit) fermentation by 1H NMR spectroscopy and chemometric analyses. J. Braz. Chem. Soc. 2012, 23, 1815−1822. (9) Fortes, G. A. C.; Naves, S. S.; Godoi, F. F. F.; Duarte, A. R.; Ferri, P. H.; Santos, S. C. Assessment of a Maturity Index in Jabuticaba Fruit by the Evaluation of Phenolic Compounds, Essential Oil Components, Sugar Content and Total Acidity. Am. J. Food Technol. 2011, 6, 974− 984. (10) Magallhães, M. M.; Barros, R. S.; Finger, F. L. Changes of structural carbohydrates in developing fruit of Myrciaria jaboticaba. Sci. Hortic. 1996, 66, 17−22. (11) Lepš, J.; Šmilauer, P. Multivariate Analysis of Ecological Data Using Canoco; Cambridge University Press: Cambridge, UK, 2007; 269 pp. (12) Ward, J. H. Hierarchical grouping to optimize an objective function. J. Am. Stat. Assoc. 1963, 58, 238−244. (13) Borcard, D.; Legendre, P.; Drapeau, P. Partialling out the spatial component of ecological variation. Ecology 1992, 73, 1045−1055. J


Article

Journal of Agricultural and Food Chemistry at different stages of berry development. Span. J. Agric. Res. 2010, 8, 425−433. (33) Artés-Hernández, F.; Artés, F.; Allende, A. Sugar composition changes in ‘Autumn Seedless’ table grape during long term cold storage. Acta Hortic. 2003, 628, 363−366. (34) Jham, G. N.; Fernandes, S. A.; Garcia, C. F.; Palmquist, D. Comparison of GC and HPLC for quantification of organic acids in two jaboticaba (Myrciaria) fruit varieties. Quim. Nova 2007, 30, 1529− 1534. (35) Davey, M. W.; Montagu, M.; Inze, D.; Sanmartin, M.; Kanellis, A.; Smirnoff, N. Plant L-ascorbic: chemistry, function, metabolism, bioavailable and effects of processing. J. Sci. Food Agric. 2000, 80, 825− 860. (36) Reynertson, K. A.; Yang, H.; Jiang, B.; Basile, M. J.; Kennelly, E. J. Quantitative analysis of antiradical phenolic constituents from fourteen edible Myrtaceae fruits. Food Chem. 2008, 109, 883−890. (37) Treutter, D. Managing Phenol Contents in Crop Plants by Phytochemical Farming and BreedingVisions and Constraints. Int. J. Mol. Sci. 2010, 11, 807−857. (38) Wang, S. Y.; Zheng, W. Effect of plant growth temperature on antioxidant capacity in strawberry. J. Agric. Food Chem. 2001, 49, 4977−4982. (39) Tuberoso, C. I. G.; Melis, M.; Angioni, A.; Pala, M.; Cabras, P. Myrtle hydroalcoholic extracts obtained from different selections of Myrtus communis L. Food Chem. 2007, 101, 806−811. (40) Quideau, S. Chemistry and biology of ellagitannins-An Underestimated Class of Bioactive Plant Polyphenols; World Scientific Publishing Co. Pte. Ltd.: Singapore, 2009; 374 pp. (41) Salminen, J.-P.; Roslin, T.; Karonen, M.; Sinkkonen, J.; Pihlaja, K.; Pulkkinen, P. Seasonal variation in the content of hydrolyzable tannins, flavonoid glycosides, and proanthocyanidins in oak leaves. J. Chem. Ecol. 2004, 30, 1693−1711. (42) Fadda, A.; Mulas, M. Chemical changes during myrtle (Myrtus communis L.) fruit development and ripening. Sci. Hortic. 2010, 125, 477−485. (43) Del Bubba, M.; Giordani, E.; Pippucci, L.; Cincinelli, A.; Checchini, L.; Galvan, P. Changes in tannins, ascorbic acid and sugar content in astringent persimmons during on-tree growth and ripening and in response to different postharvest treatments. J. Food Compos. Anal. 2009, 22, 668−677. (44) García-Villalba, R.; Espín, J. C.; Aaby, K.; Alasalvar, C.; Heinonen, M.; Jacobs, G.; Voorspoels, S.; Koivumäki, T.; Kroon, P. A.; Pelvan, E.; Saha, S.; Tomás-Barberán, F. A. Validated Method for the Characterization and Quantification of Extractable and Nonextractable Ellagitannins after Acid Hydrolysis in Pomegranate Fruits, Juices, and Extracts. J. Agric. Food Chem. 2015, 63, 6555−6566. (45) Leite-Legatti, A. V.; Batista, Â . G.; Dragano, N. R. V.; Marques, A. C.; Malta, L. G.; Riccio, M. F.; Eberlin, M. N.; Machado, A. R.T.; Silva, L. B. D.; Ruiz, A. L. T. G.; Carvalho, J. E.; Pastore, G. M.; Maróstica, M. R., Jr Jaboticaba peel: Antioxidant compounds, antiproliferative and antimutagenic activities. Food Res. Int. 2012, 49, 596−603. (46) Montes, C.; Vicario, I. M.; Raymundo, M.; Fett, R.; Heredia, F. J. Application of tristimulus colorimetry to optimize the extraction of anthocyanins from Jaboticaba (Myricia Jaboticaba Berg.). Food Res. Int. 2005, 38, 983−988. (47) Fracassetti, D.; Costa, C.; Moulay, L.; Tomás-Barberán, F. A. Ellagic acid derivatives, ellagitannins, proanthocyanidins and other phenolics, vitamin C and antioxidant capacity of two powder products from camu-camu fruit (Myrciaria dubia). Food Chem. 2013, 139, 578− 588. (48) Glabasnia, A.; Hofmann, T. Sensory directed identification of taste-active ellagitannins in American (Quercus alba L.) and European oak wood (Quercus robur L.) and quantitative analysis in bourbon whiskey and oak-matured red wines. J. Agric. Food Chem. 2006, 54, 3380−3390. (49) Chassaing, S.; Lefeuvre, D.; Jacquet, R.; Jourdes, M.; Ducasse, L.; Galland, S.; Grelard, A.; Saucier, C.; Teissedre, P. L.; Dangles, O.; Quideau, S. Physicochemical Studies of New Anthocyano-Ellagitannin

Hybrid Pigments: About the Origin of the Influence of Oak CGlycosidic Ellagitannins on Wine Color. Eur. J. Org. Chem. 2010, 2010, 55−63. (50) Matsuo, Y.; Wakamatsu, H.; Omar, M.; Tanaka, T. Reinvestigation of the Stereochemistry of the C-Glycosidic Ellagitannins, Vescalagin and Castalagin. Org. Lett. 2015, 17, 46−49. (51) Agrawal, A. A.; Hastings, A. P.; Johnson, M. T. J.; Maron, J. L.; Salminen, J.-P. Insect Herbivores Drive Real- Time Ecological and Evolutionary Change in Plant Populations. Science 2012, 338 (6103), 113−116. (52) Fait, A.; Hanhineva, K.; Beleggia, R.; Dai, N.; Rogachev, I.; Nikiforova, V. J.; Fernie, A. R.; Aharoni, A. Reconfiguration of the Achene and Receptacle Metabolic Networks during Strawberry Fruit Development. Plant Physiol. 2008, 148, 730−750. (53) Conde, C.; Silva, P.; Fontes, N.; Dias, A. C. P.; Tavares, R. M.; Sousa, M. J.; Agasse, A.; Delrot, S.; Gerós, H. Biochemical Changes throughout Grape Berry Development and Fruit and Wine Quality. Food 2007, 1, 1−22.

K


Polyphenol and Ellagitannin Constituents of Jabuticaba (Myrciaria

Recommend Documents