Structural and Biochemical Changes Induced by Pulsed Electric Field

Article pubs.acs.org/JAFC

Structural and Biochemical Changes Induced by Pulsed Electric Field Treatments on Cabernet Sauvignon Grape Berry Skins: Impact on Cell Wall Total Tannins and Polysaccharides Céline Cholet,†,§ Cristèle Delsart,†,§ Mélina Petrel,‡,⊥,# Etienne Gontier,‡,⊥,# Nabil Grimi,⊗ Annie L’Hyvernay,†,§ Remy Ghidossi,†,§ Eugène Vorobiev,⊗ Martine Mietton-Peuchot,†,§ and Laurence Gény*,†,§ †

Université de Bordeaux, Institut des Sciences de la Vigne et du Vin, EA 4577, Unité de recherche œnologie, France INRA, ISVV, USC 1219 Œnologie, 210 Chemin de Leysotte, CS 50008, F-33882 Villenave d’Ornon, France ‡ Université Bordeaux, Bordeaux Imaging Center, UMS 3420, F-33000 Bordeaux, France ⊥ CNRS, Bordeaux Imaging Center, UMS 3420, F-33000 Bordeaux, France # INSERM, Bordeaux Imaging Center, US 004, F-33000 Bordeaux, France ⊗ Université de Technologie de Compiègne, Transformation Intégrée de la Matière Renouvelable (TIMR, EA 4297, UTC/ESCOM), Equipe Technologies Agro-industrielles, Centre de Recherche de Royallieu, B.P. 20529-60205, Compiègne Cedex, France §

ABSTRACT: Pulsed electric field (PEF) treatment is an emerging technology that is arousing increasing interest in vinification processes for its ability to enhance polyphenol extraction performance. The aim of this study was to investigate the effects of PEF treatment on grape skin histocytological structures and on the organization of skin cell wall polysaccharides and tannins, which, until now, have been little investigated. This study relates to the effects of two PEF treatments on harvested Cabernet Sauvignon berries: PEF1 (medium strength (4 kV/cm); short duration (1 ms)) and PEF2 (low intensity (0.7 kV/cm); longer duration (200 ms)). Histocytological observations and the study of levels of polysaccharidic fractions and total amounts of tannins allowed differentiation between the two treatments. Whereas PEF1 had little effect on the polyphenol structure and pectic fraction, PEF2 profoundly modified the organization of skin cell walls. Depending on the PEF parameters, cell wall structure was differently affected, providing variable performance in terms of polyphenol extraction and wine quality. KEYWORDS: microscopy, polysaccharides, pulsed electric field, cell wall tannins, Vitis vinifera L.

■

structure.2,3,6,7,19 A first study has been carried out linking the biochemical study of must and wine with a study of the histological structure of grape berries immediately after two PEF treatments.19 The histological structure of the pericarp, or carpellary wall, of the grape berry is composed of three distinct tissues: the skin or exocarp including the cuticle, the epidermis, and the hypodermis; the mesocarp or pulp made up of large cells; and the endocarp comprising a thin layer of inner epidermis cells in contact with the seeds.21−24 Throughout berry growth, internal parenchyma cells of the mesocarp and inner hypodermal cells become pulp cells: vacuolar phenolic compounds decrease and cell walls become progressively thinner from the inside toward the outside of the pericarp.25−27 The skin of mature berries is thinned; this corresponds to the external epidermis with a thick cuticle and a thin external hypodermis. Skin cell walls are always thick. External epidermal cells are frequently senescent, and the majority of them contain no vacuolar phenolic compounds. The majority of external hypodermis cells still contain vacuolar phenolic compounds.28 The morphological

INTRODUCTION The application of various pulsed electric field (PEF) treatments to living cells induced electrical electroporation phenomena on the cell membranes.1−4 These permeabilizations of cell membranes are due to the formation of pores of various sizes.4 This phenomenon is reversible or not, depending on the parameters of the PEF,1,3 in particular the intensity and duration of the treatment. In fact, every living cell has a natural capacity for resistance to external transmembrane potential to which it is subjected, up to a limit called the “critical value”. Beyond this limit electroporation becomes irreversible. Thus, the greater the strength of PEF beyond the critical value of the treated cell, the more intense and irreversible is the phenomenon of electroporation.5 For microorganisms, the strength of the electric field required to exceed the critical value lies somewhere between 5 and 55 kV/cm.6−11 For plant cells of larger volume, it lies between 0.1 and 10 kV/cm.12,13 Thus, with high-strength PEF treatment, the electroporation mechanism induced the destruction of cell membrane structures11 and seemed to lead to cytological disruption of a large portion of yeast cells.6,8 PEF treatment of medium strength promoted the extraction of cellular components in plant cells,14 and it is this phenomenon that is arousing more and more interest in the world of winemaking.15−20 To date, very few studies have investigated the effect of PEF treatment on histocytological © 2014 American Chemical Society

Received: Revised: Accepted: Published: 2925

October 24, 2013 February 16, 2014 March 11, 2014 March 11, 2014 dx.doi.org/10.1021/jf404804d | J. Agric. Food Chem. 2014, 62, 2925−2934

Journal of Agricultural and Food Chemistry

Article

for the treatment. Each series consisted of n pulses, pulse duration (ti), time interval between pulses (Δt), and there was a pause (Δtt) after each train of pulses. The effective time of the PEF treatment was regulated by the variation of the number of series N and is calculated as tPEF = nNti. A sample of grape berries (0.7−4.2 kg) was placed in the PEF treatment cell between two stainless plane electrodes (230 mm × 265 mm). The distance between electrodes was 70 mm for PEF1 and 12 mm for PEF2. The value of the electric field E was evaluated as E = U/d, where U is the applied voltage. The initial conductivity of the samples was approximately 2.5 mS/cm. The specific energy consumption was calculated using the following formula: W = UItPEF/m, where m is the mass of grapes. These protocols allowed fine regulation of the treatment without any noticeable temperature elevation (ΔT ≤ 6 °C for PEF1 and ΔT ≤ 14 °C for PEF2) during the PEF treatment. Three batches of 50 “control” berries were collected on the same day as the treated berries and stored in the same way to await the different biochemical analyses. Microscopy Skin Characterization and Cytological Preparation for Light and Transmission Electron Microscopy (Method Described by Colin et al.35). A first sampling of 10 berries from each batch, treated and untreated, was carried out immediately after the various PEF treatments (day 0) to observe the immediate effects of the treatment on their cellular structure. A second sampling was carried out after the end of alcoholic fermentation (day 14) to note the indirect effect of the treatment on extraction from cells. For each PEF treatment, 10 berries were randomly selected and immediately prepared for microscopy as described by McManus36 for fixation and by Thiéry37 for staining. Control (untreated) and treated berries were dissected into four sections each, always in the same manner and fixed (4 °C for 90 min in 2.5% glutaraldehyde in a phosphate buffer, pH 7.2), then washed in the same buffer, and postfixed (4 °C for 90 min in osmium tetroxide). The berry pieces were dehydrated progressively in successive baths of ethyl alcohol from 40% (v/v) to absolute (100% v/v) and were then embedded in epoxy resin (Epon). Light Microscopy (MO). Semithin sections (2 μm thick) were stained via the periodic acid Schiff (PAS) method for the detection of polysaccharides.37 Two hundred cross sections were observed by light microscopy for each treatment mode. Ten cross sections per treatment mode were photographed using an Olympus camera. The PAS-stained sections were used to determine the general histocytological morphology and to evaluate the polysaccharide content of the cell walls according to the different shades of pink obtained (negative PAS = light pink, positive PAS = dark and intense pink). Transmission Electron Microscopy (TEM). Thin sections were cut using an ultramicrotome UCT (LEICA, Vienna, Austria) and picked up on 150-mesh copper grids. The sections were counterstained with uranyl acetate and lead citrate.38 One hundred cross sections were examined for each treatment using a transmission electron microscope at 120 kV (Tecnai 12, FEI) and photographed with an ORIUS 832-11 MPixel (Gatan) camera. Biochemical Analysis of Total Flavan-3-ols and Pectic Fractions of Skin Cell Walls. Frozen berries were peeled with a razor blade to retain only their skins. The skins were then very finely ground in a mortar under liquid nitrogen until a powder form was obtained. Extractions of Cell Wall Fraction.32,39 The cell wall isolation procedure followed the method described by Harris et al.40 with slight modifications to allow us to perform the assay of tannins and polysaccharides on the same extraction sample. One and a half grams of “skin powder” was suspended in 10 mL of 0.2 M Tris-HCl buffer, pH 7.5, containing 2.5% (w/v) EDTA (homogenization buffer) and homogenized at 4 °C to prevent the activation of the pectolytic enzymes41 and the polyphenol oxidases.42 This sample was then centrifuged at 15000 rpm for 20 min at 4 °C. The resulting pellet was resuspended in 10 mL of homogenization buffer and recentrifuged at 15000 rpm for 30 min at 4 °C. The pellet was further resuspended twice in 10 mL of 2.5 M saccharose and recentrifuged at 15000 rpm for 30 min at 4 °C. The pellet was

study of tissues (histological observations) and cells (cytological observations) confirms the biochemical assays: on the one hand, during grape berry development there are textural changes in the cell walls that lead to softening of the cell walls;29 and on the other hand, green-berry tissues are rich in phenolic compounds. During berry development, phenolic compound content decreases until veraison (grape color change) and then stabilizes until maturity.30−32 Tannins in the internal cell fractions are polymerized to a lesser degree than those in the cell wall.32,33 All of these histological and cytological phenomena are essential to the extraction of both juice from the pulp and phenolic and aromatic compounds from the skin. The degree of extraction of these compounds has a significant influence on the duration of maceration and the quality of the resulting wine. Thus, the red wines of Bordeaux ought to be quite tannic to be well structured and to have good aging potential.34 Throughout maceration, tannins are extracted from either skins or seeds, but during conventional winemaking it is very difficult both to extract seed tannins and to know how much tannin is extracted from seeds. Therefore, the availability of a technique which enhances the extraction of tannins is advantageous in that such a technique may produce a better tannin balance. Moreover, in the case of harvests that are not fully ripe, maceration ought to be of longer duration to improve the extraction rate. However, this may concomittantly release unwanted flavors (vegetable aromas, for example). Thus, the use of a technique that reduces maceration time may avoid such complications. According to recent studies, the development of an extraction process such as PEF technology in the wine industry seems to be one of the less time-consuming and more energy-efficient18,20 alternatives. Indeed, this method has proved highly efficient in terms of both molecule extraction and yield.15−20 In previous works, we studied the effects of PEF treatment on must and wine composition.19,20 The present work is complementary to our previous study and shows the effects of PEF treatment on the histocytological structure and the cell wall composition of berry, providing a better understanding of how polyphenol extraction can be modified by PEF. The objective was to study the effects on grape berries of two pulsed field treatments of medium strength. Light and transmission electron microscopy were both used to observe structural changes to the skin cell walls of treated grape berries. These observations completed the results obtained in biochemical analyses of skin cell walls of the grape berries examined.

■

MATERIALS AND METHODS

Plant Material. One hundred and twenty kilograms of grapes berries of Vitis vinifera L. cv. Cabernet Sauvignon were manually harvested at maturity (sugars = 225 g/L; extractability of anthocyanins = 50%; total polyphenols richness = 48) in a Bordeaux vineyard in September 2009. These berries were then destemmed by hand. PEF Treatments. Three batches of 20 kg of lightly crushed and pressed berries were prepared, two of which were subjected to two different PEF treatments. The third batch corresponded to the control group. The PEF parameters described below were chosen in to compare the effects of a treatment of short duration (1 ms) and high strength (4 kV/cm) with the effects of a treatment of long duration (200 ms) and low strength (0.7 kV/cm). Cabernet Sauvignon grape berries were placed between two parallel stainless steel electrodes, and PEF was subsequently applied. [PEF1: E = 4 kV/cm; tPEF = 1 ms, W = 3.99 Wh/kg (n = 1; ti = 10 μs; Δt = 10 ms; Δtt = 10 s; N = 100). PEF2: E = 0.7 kV/cm; tPEF = 200 ms; W = 31 Wh/kg (n = 100; ti = 100 μs; Δt = 10 ms; Δtt = 10 s; N = 20).] N series of pulses were used 2926

dx.doi.org/10.1021/jf404804d | J. Agric. Food Chem. 2014, 62, 2925−2934


Article

resuspended six more times in 10 mL of homogenization buffer and centrifuged anew after each resuspension, each time with different settings: 16000 rpm, 30 min, 4 °C; 14000 rpm, 30 min, 4 °C; 10000 rpm, 20 min, 4 °C; 9000 rpm, 15 min, 4 °C; 8000 rpm, 10 min, 4 °C; 5000 rpm, 10 min, 4 °C. The pellet was resuspended in 10 mL of Triton X-100 0.1% and recentrifuged at 3000 rpm for 10 min at 4 °C. The pellet was resuspended twice in 10 mL of homogenization buffer and recentrifuged at 3000 rpm for 10 min at 4 °C and at 2000 rpm for 5 min at 4 °C. The sample was subsequently filtered through a 3 μm PTFE filter and dried in an oven at 35 °C. This fraction was designated the “cell wall fraction of the skin”. Extraction of Polysaccharidic Fractions of the Cell Wall Fraction of the Skin (Adapted from Saulnier and Thibault 43). For all samples, extraction and analysis were performed in triplicate. One-tenth of a gram of the cell wall fraction of the skin was suspended in a mixture of 20 μL of ethanol 95% and 20 mL of distilled water, stirred for 24 h at room temperature, and centrifuged at 1200 rpm for 30 min. The resulting supernatant corresponded to the PSE fraction (polysaccharides soluble in water). The pellet was resuspended in 20 mL of ammonium oxalate 2% (w/p), stirred for 2 h at room temperature, and centrifuged at 1200 rpm for 30 min. The resulting supernatant corresponded to the PSOX fraction (polysaccharides soluble in ammonium oxalate). The pellet was again suspended in 20 mL of 0.05 N HCl, stirred for 2 h at 85 °C, and centrifuged at 1200 rpm for 30 min. The resulting supernatant corresponded to the PSH fraction (polysaccharides soluble in acid). The pellet was once more suspended in 20 mL of 0.05 N NaOH, stirred for 2 h at 4 °C, and centrifuged at 1200 rpm for 30 min. The resulting supernatant corresponded to the PSOH fraction (polysaccharides soluble in sodium hydroxide). The pellet corresponded to the residual matter and was designated RP (residual pellet). Extraction of Tannins.44 For all samples, extraction was performed in triplicate. Either 0.1 g of the cell wall fraction of the skin and the residual pellet or 10 mL of pectic fraction was suspended in 10 mL of methanol containing 0.1% HCl. This solution was stirred for 3 h at 20 °C. The extract was filtered through 20 μm PTFE filters. For pectic fractions, this extract was designated the “phenolic extract of the skin pectic fraction”. In the same manner as for the cell wall fraction of the skin and the residual pellet, the powder remaining on the filter was suspended in 10 mL of methanol containing 0.1% HCl (v/v) and stirred anew for 3 h at 20 °C. The resulting extract was filtered through 20 μm PTFE filters. This extract was designated the “phenolic extract of skin cell wall” from the cell wall fraction or the “phenolic extract of the residual pellet” from the residual pellet. Analysis of Tannins by Reverse Phase HPLC following Phloroglucinolysis.33,34,45 The mean degree of polymerization (mDP) and the subunit composition of procyanidins in the different fractions were analyzed. Six hundred microliters of phenolic extract, desiccated under nitrogen flow, plus 200 μL of reagent ( 0.2 Nmethanol-HCl + ascorbic acid + phloroglucinol) were mixed and incubated at 50 °C for 20 min. The reaction was stopped with 200 μL of sodium acetate. This extract was filtered and placed in a 2 mL HPLC sealed vial. Ten microliters was injected into the HPLC for analysis (column, 250 × 4.6 μm, 5 μm, ODS (Beckman, Roissy Charles de Gaulle, France): precolumn, 10 × 4.6 mm, 5 μm, BDS C18 (Thermo Hypersil); flow rate, 1 mL/min; solvent A, water/acetic acid (19:1 v/v); solvent B, MeOH/acetic acid (19:1 v/v); gradient, 5% B from 0 to 30 min, 20% B from 30 to 55 min, 40% B from 55 to 60 min, 90% B from 60 to75 min, 5% B from 75 to 80 min; injection volume, 10 μL; detection wavelength, 280 nm. Spectrophotometric Analysis of Polysaccharides.46 One hundred and fifty microliters of extract diluted at 1/10 plus 750 μL of 95% H2SO4 were stirred for 3 min at 0 °C and then for 6 min at 100 °C and then cooled for 10 min at 0 °C. After the addition of 15 μL of 3hydroxydiphenyl (MHDP), the extracts were left in the dark for 20 min at room temperature before dosing at a wavelength of 520 nm. Statistical Data Treatment. Two-way analysis of variance (ANOVA) could not be performed (non-normality of the residues and/or nonhomogeneity of the variance). We thus performed nonparametric tests. The statistical significance of differences was

determined by the Freidman test (α = 0.05) for paired samples and by the Kruskal and Wallis test (α = 0.05) for unpaired samples. Experimental data detected as being significantly different are marked in the tables with different letters.

■

RESULTS Immediate and Long-Term Effects of Different Pulsed Electric Field Treatments on Skin Histocytological Structure. Skin histocytological structures of grape berries were observed in both light microscopy (200 cross sections) and transmission electron microscopy (100 cross sections) for the two PEF treatment modes. Observations were carried out soon after treatment (day 0) to observe the immediate effects of PEF treatment and at the end of the alcoholic fermentation (day 14) to note the modifications to the cell structure of the grape berry skin.

Figure 1. Evolution of the skin cross sections immediately after PEF treatments (day 0) (A, untreated (0 kV/cm); B, PEF1-treated (4 kV/ cm; 1 ms; 4 Wh/kg); C, PEF2-treated (0.7 kV/cm; 200 ms; 31 Wh/ kg)) and 14 days after PEF treatments (day 14) (A′, untreated (0 kV/ cm); B′, PEF1-treated (4 kV/cm; 1 ms; 4 Wh/kg); C′, PEF2-treated (0.7 kV/cm; 200 ms; 31 Wh/kg)), under optical microscopy. eE, external epidermis; oH, outer hypodermis; iH, inner hypodermis; S, skin; VPC, vacuolar phenolic compounds.

Control Berries (Figures 1, 2, and 4). At day 0, all berry skins observed were characterized by one layer of epidermal cells and by five to six layers of hypodermal cells (Figure 1A), which were found to be rich in organelles (o) such as mitochondria (m) (Figure 2A). Vacuolar polyphenols (VPC) appeared as dense amorphous precipitates from the epidermis and the outer hypodermis (Figures 1A and 2A) and as gray granules originating from the inner hypodermis (Figures 1A and 2B). Despite slight staining with PAS (Figure 1A), it was 2927



Article

the cell walls of the outer hypodermis showed perforations (Figure 4A′, arrowed). At the inner hypodermis, fiber cell wall polysaccharides were more distended than in the control batch (Figure 4, panel B vs panel B′).

Figure 2. Skin cross sections of (A, B, C) untreated-berries (0 kV/cm) and of (A′, B′, C′) PEF1-treated berries (4 kV/cm; 1 ms, 4 Wh/kg) observed at day 0, under transmission electron microscopy: (A, A′) outer hypodermis; (B, B′) inner hypodermis; (C, C′) outer hypodermis. CW, cell wall; o, organelles; ml, middle lamella; m, mitochondria; VPC, vacuolar phenolic compounds. Figure 3. Skin cross sections of (A, B, C) untreated berries (0 kV/cm) and of (A*, B*, C*) PEF2-treated berries (0.7 kV/cm; 200 ms, 31 Wh/kg) observed at day 0, under transmission electron microscopy: (A, A*) outer hypodermis; (B, B*) inner hypodermis; (C, C*) outer hypodermis. CW, cell wall; iH, inner hypodermis; m, mitochondria; ml, middle lamella; o, organelles; oH, outer hypodermis; p, plastid; VPC, vacuolar phenolic compounds.

possible to observe that the cell walls (CW) showed no signs of deterioration (Figure 2A−C). At 14 days, almost all of the vacuolar polyphenols had been evacuated (Figure 1A′). Very few vacuolar polyphenols were still present in the epidermis and outer hypodermis, where they appeared as very dense, dark red or brown, amorphous precipitates. Curiously, contrary to day 0, cell walls were found to show homogeneous dark pink PAS staining (Figure 1A′), and the contrast under TEM was seen to be homogeneous and electron-dense (Figure 4A,B). Most organelles, such as plastids (p), were senescent (Figure 4B). PEF 1 (4 kV/cm, 1 ms, 4 Wh/kg) (Figures 1, 2, and 4). At day 0, the skin of all treated berries was more distended (Figure 1B) than that of the control batch (Figure 1A). Vacuolar polyphenols of the epidermis (Figure 2A′) and hypodermis (Figure 2B′) were less dispersed and more condensed in the vacuole. Small spherical structures, which were localized only at the middle lamella level in the control batch (Figure 2C′), were observed over a large portion of the hypodermis cell wall thickness (Figure 2C′). At day 14 (Figure 1B′), the hypodermis of all treated berry skins contained no vacuolar phenolic compounds with a dense dark red or brown aspect, as was the case in the control batch. However, it was richer in vacuolar phenolic compounds in the form of small gray granules dispersed throughout the vacuolar space. It can therefore be concluded that these phenolic compounds were probably chemically different from those present in the control berries at the same time. Furthermore,

PEF 2 (0.7 kV/cm, 200 ms, 31 Wh/kg) (Figures 1, 3, and 4). At day 0, in contrast to PEF1, the skin of all treated berries was seen to be much more degraded than in the control batch, especially in the deepest layers of the inner hypodermis of the skin (Figure 1C). The vacuolar polyphenols of the outer cell layers of the skin appeared denser and more contrasted, as was the case for PEF1. The outer hypodermis cells (Figure 3A*), and even more so those of the inner hypodermis (Figure 3B*), were disorganized, showing destructuring of the middle lamella of cell walls (arrowed) and organelles, such as mitochondria (m) or plastids (p), that had been disrupted. At highest magnification, the aspect of the cell wall appeared unstructured, with the presence of small spherical structures throughout the whole thickness (Figure 3C*). At day 14 the skin of the treated berries was, in 80% of cases, less rich in vacuolar polyphenols than that of the untreated berries. However, some areas could be observed in which phenolic compounds of a dense gray appearance persisted in the cells (Figure 1C′). In all cases, cell wall structure and the organization of the polysaccharide fibers had a very different 2928



Article

Tangential walls of contiguous layers of cells were asymmetrical on either side of the middle lamella (Figure 4B*). Moreover, the middle lamella was observed under TEM as being more highly contrasted and electron dense (Figure 4B*), which could be one of the consequences of changes in the wall that contained more sites where the counterstaining metal salts of the different sections could be lodged. In conclusion, the immediate impact of PEF1 was characterized by a distension of the skin and a condensation of most of the vacuolar polyphenols. The impact on the cell walls was more limited, but remained significant, at 14 days after perforations were first observed. The immediate impact of PEF2 was much more pronounced: the skin became very dense, the polyphenols were more highly concentrated, and, more especially, the cell walls were greatly modified. They were observed at 14 days as being very eroded and degraded. Immediate Effects of Pulsed Electric Field Treatments on Skin Cell Wall Condensed Tannins. PSE and PSOX fractions correspond to the easily extractable peripheral pectic fractions, whereas PSH and PSOH fractions correspond to the internal pectic fractions that are only extractable with greater difficulty.42 The residual pellet (RP) corresponds to the “heart” of the cell wall. Control Berries (Table 1). The pectic fractions richest in condensed tannins were PSH and PSOH, which were internal and strongly bonded with the cell wall fibers. The larger condensed tannins with the highest mDP were found in the PSH fraction. PEF 1 (4 kV/cm, 1 ms, 4 Wh/kg) (Table 1). In the different peripheral cell wall fractions (PSE; PSOX), the treatment did not have a marked effect. However, levels of condensed tannins in the PSE fraction (TPSE) tended to be higher (C, 0.17, vs PEF1, 0.24) and mDP tended to decrease (C, 3.42, vs PEF1, 2.91), reflecting a probable decondensation of tannins and their accumulation in this fraction. In the PSOX fraction, tannin levels (TPSOX) tended to be lower (C, 0.29, vs PEF1, 0.2) and mDP tended to increase (C, 2.43, vs PEF1, 3.26), reflecting a probable loss of tannins further to their condensation in this fraction.32 In the different internal cell wall fractions (PSOH,

Figure 4. Skin cross sections of (A, B) untreated berries (0 kV/cm), (A′, B′) PEF1-treated berries (4 kV/cm; 1 ms, 4 Wh/kg), and (A*, B*) PEF2-treated berries (0.7 kV/cm; 200 ms, 31 Wh/kg) observed at day 14, under transmission electron microscopy: (A, A′, A*, B*) outer hypodermis; (B, B′) inner hypodermis. CW, cell wall; eE, external epidermis; iH, inner hypodermis; ml, middle lamella; oH, outer hypodermis; p, plastid; VPC, vacuolar phenolic compounds.

appearance from the control batch. In treated skins, cell angles were observed to be rather distended with high-contrast deposits and perforations that evoke meatus (Figure 4A*).

Table 1. Effects of PEF1 and PEF2 on the Total Tannins and mDP of the PSE, PSOX, PSH, PSOH, and RP Polysaccharide Fractions in Berry Skinsa C

PEF1 (4 kV/cm, 1 ms, 4 Wh/kg)

PEF2 (0.7 kV/cm, 200 ms, 31 Wh/kg)

TPSE

tannins of polysaccharide fractions total tannins (mg/g of skin) mDP

0.17 ± 0.11 a* 3.42 ± 0.88 a*

0.24 ± 0.17 ab* 2.91 ± 0.49 a*

0.42 ± 0.04 b* 3.17 ± 2.78 a*

TPSOX

total tannins (mg/g of skin) mDP

0.29 ± 0.08 a* 2.43 ± 0.50 a*

0.20 ± 0.03 a* 3.26 ± 0.72 a*

0.14 ± 0.05 b* 2.79 ± 1.13 a*

TPSH


2.81 ± 0.83 a* 15.83 ± 5.07 a*

1.13 ± 1.36 a* 58.03 ± 68.8 a*4

2.17 ± 1.59 a* 13.84 ± 1.22 a*

TPSOH


2.57 ± 1.85 a* 3.59 ± 3.75 a*

5.19 ± 5.49 a* 1.63 ± 0.25 a*

9.29 ± 8.03 b* 1.64 ± 0.13 a*

TRP


0.75 ± 0.15 a* 5.79 ± 2.11 a*

0.66 ± 0.64 a* 2.64 ± 0.32 a*

1.67 ± 0.87 a* 4.79 ± 1.99 a*

a Values are means ± standard deviations. (*) Significant difference in Kruskal and Wallis test (α = 0.1). PEF1, treatment 1 of pulsed electric field; PEF2, treatment 2 of pulsed electric field; C, control berries; TPSE, tannins of PSE; TSPOX, tannins of PSOX; TPSH, tannins of PSH; TPSOH, tannins of PSOH; TRP, tannins of RP; PSE, extracted polysaccharide fraction in water; PSOX, extracted polysaccharide fraction in oxalate; PSH, extracted polysaccharide fraction in acid; PSOH, extracted polysaccharide fraction in sodium hydroxide; RP, pellet of residual polysaccharide fractions.

2929



Article

PSH, RP), the same phenomenon was observed because levels of condensed PSH tannins (TPSH) tended to be lower (C, 2.81, vs 1.13), and mDP levels tended to increase (C, 15.83, vs PEF1, 58.03), as was the case for the PSE fraction. The tannin level of the PSOH fraction (TPSOH) tended to be higher (C, 2.57, vs PEF1, 5.19), and mDP tended to decrease (C, 3.59, vs PEF1, 1.63), as was the case for the PSOX fraction. Moreover, in the residual pellet (TRP), mDP tended to decrease (C, 5.79, vs PEF1, 2.64), whereas levels of condensed tannins showed no variation (C, 0.75, vs PEF1, 0.66). This could reflect a breakdown of the tannins present in this fraction and a leakage of decondensed tannins into adjacent fractions. PEF2 (0.7 kV/cm, 200 ms, 31 Wh/kg) (Table 1). The two peripheral cell walls (PSE; PSOX) were affected by the treatment as compared to the control batch. The level of condensed PSE tannins (TPSE) increased significantly (C, 0.17, vs PEF2, 0.42) and mDP also tended to increase (C, 3.42, vs PEF2, 4.26). In the PSOX fraction the level of condensed tannins (TPSOX) decreased significantly (C, 0.29, vs PEF2, 0.14) and mDP also tended to increase (C, 2.43, vs PEF2, 2.79). It is conceivable that either the TPSOX were depleted after their condensation in this fraction or a part of TPSOX was decondensed under the action of PEF2, which spontaneously migrated from this fraction to accumulate in PSE, in which bonds are of the same nature, and spontaneously condensed in this fraction where the amount of condensed tannins increased. This migration would thus explain the large increase in the level of TPSE, whereas mDP also showed an increase. The PSE fraction was enriched in condensed tannins from TPSE and TPSOX (therefore, the average value of mDP was higher in these fractions), contributing to the significant decrease in the level of TPSOX. Among the internal fractions (PSOH; PSH; RP) only PSOH was significantly affected; mDP tended to decrease (C, 3.59, vs PEF2, 1.64), reflecting a rupture due to treatment, and the level of condensed tannins (TPSOH) increased (C, 2.57, vs PEF2, 9.29), reflecting an accumulation of these tannins. Moreover, the mDP of condensed PSH tannins (TPSH) tended to be lower than in the control batch (C, 15.83, vs PEF2, 13.84), reflecting a rupture, whereas the level of TPSH tended to decrease (C, 2.81, vs PEF2, 2.17). This decrease in the level of TPSH can be explained by a leakage of TPSH to the PSOH fraction, contributing to the increased levels of TPSOH. In the RP, mDP tended to decrease as compared with the control batch (C, 5.79, vs PEF2, 4.79) and the level of condensed tannins (TRP) increased (C, 0.75, vs PEF2, 1.67), reflecting an accumulation of these decondensed tannins. Finally, whereas PEF1 did not significantly affect the tannic skeleton of skin cell walls, the consequences of PEF2 were much more significant: the tannic skeleton of skin cell walls was highly disorganized, especially the external fractions. Immediate Effects of Different Pulsed Electric Field Treatments on Skin Cell Wall Polysaccharidic Fractions. The peripheral pectic fractions PSE and PSOX are easily extractable because they are connected by weak bonds. The internal pectic fractions PSH and PSOH are not easily extractable because they are connected by strong bonds. Each of the fractions contains mainly galacturonic acid, arabinose, and galactose, which correspond to pectic compounds.43 It is thus possible to refer indifferently to the polysaccharide fraction or the pectic fraction. Control Berries (Table 2). In descending order, the most abundant polysaccharide fraction corresponded to the PSE

Table 2. Effects of PEF1 and PEF2 on the Contents of External Polysaccharide Fractions (PSE, PSOX) and Internal Polysaccharide Fractions (PSH, PSOH) in Berry Skinsa polysaccharide content (mg/g of skin) PSE fractions PSOX fractions PSH fractions PSOH fractions

PEF1 (4 kV/cm, 1 ms, 4 Wh/kg)

C 8.80 1.32 0.70 2.59

± ± ± ±

2.25 0.36 0.07 0.50

a* a* a* a*

9.71 1.57 0.56 1.76

± ± ± ±

3.12 0.47 0.07 0.74

a* a* b* a*

PEF2 (0.7 kV/cm, 200 ms, 31 Wh/kg) 3.88 2.54 0.55 1.55

± ± ± ±

2.66 0.73 0.16 0.63

b* b* c* b*

a Values are means ± standard deviations. (*) Significant difference in Kruskal and Wallis test (α = 0.1). PEF1, treatment 1 of pulsed electric field; PEF2, treatment 2 of pulsed electric field; C, control berries; PSE, extracted polysaccharide fraction in water; PSOX, extracted polysaccharide fraction in oxalate; PSH, extracted polysaccharide fraction in acid; PSOH, extracted polysaccharide fraction in sodium hydroxide.

fraction, which is the most easily extractable, followed by PSOH, PSOX, and PSH. PEF 1 (4 kV/cm, 1 ms, 4 Wh/kg) (Table 2). This treatment mode did not cause changes in the distribution of pectic fractions in the cell walls. Furthermore, external fractions of cell wall were not significantly affected (Table 2) because the levels of PSE and PSOX remained stable, but these fractions tended to increase, which seems to imply a slightly increased density of weak bonds (noncovalent) in the cell wall. However, PEF1 significantly modified the internal fractions, showing decreased levels of PSH, which is the cause of disruption of strong bonds (covalent) and a rearrangement of molecular bonds at this level. PEF2 (0.7 kV/cm, 200 ms, 31 Wh/kg) (Table 2). This treatment mode significantly affected all polysaccharidic fractions of the cell walls, principally in the peripheral fractions. At the periphery of the cell wall, the PSOX level was increased significantly and may have been enriched by some of the PSE, the level of which decreased (Table 2). However, the density of weak bonds (noncovalent) decreased in PSE and increased in PSOX; therefore, the molecular affinity of these fractions was totally modified. Toward the inside of the cell wall, as was the case for PEF1 but more pronounced, levels of PSH and PSOH were decreased significantly (Table 2), which is the cause of a decrease of strong bonds (covalent) and of an upheaval and rearrangement of molecular bonds at this level. As seen above in the case of the tannic skeleton, PEF1 had only a slight impact on the pectic skeleton of skin cell walls. Conversely, the consequences of PEF2 were more far-reaching seeing as this treatment mode caused the disorganization of cell wall polysaccharides and molecular bonds, which induced a modification of polyphenol extraction phenomenon.47−49

■

DISCUSSION The main action of PEF is the induction of a phenomenon of electroporation in the form of hydrophilic pores on small portions of cell membranes of the cells facing the electrodes. Cells exposed to the transmembrane energy of an external electric field undergo a reorganization of membrane phospholipids.4,5 In this way, the shape and size of the affected cells directly influences the phenomenon of electroporation. Grape berries being composed of heterogeneous tissues, with cell shape and membrane potential variables,21−27 the spread and consequences of PEF are not homogeneous in the skin. 2930



Article

of the PSH pectic fraction. Renard et al.50demonstrated that affinity and binding between cell wall procyanidins and polysaccharides in apples increased with the degree of polymerization of these phenolic compounds. Thus, in the present case, this polymerization may help strengthen cell walls32 and slow extraction phenomena.51,52 This would explain why, at day 14, the hypodermis of treated berries was still rich in vacuolar phenolic compounds (Figure 1). However, TPSOX, TPSH, and TRP were highly condensed by the PEF1 treatment, inducing, on the one hand, a strong bond with pectic compounds, particularly in the PSOX fraction where the number of noncovalent bonds probably increased, thus limiting their depolymerization.53 On the other hand, the formation of new spaces in the heart of the cell walls resulted in a diminished extractability of these tannins, which are more strongly encapsulated within microfibrillar pores and therefore strongly adsorbed by cell wall components.47 Thus, PEF1 slowed the phenomena of diffusion and extraction of most tannins from the skin of treated berries. It was also noted that the cytological aspect of vacuolar phenolic compounds was modified after the PEF1 treatment: they appeared more condensed in the cell vacuoles. This seems to demonstrate that the PEF1 treatment had had an effect mainly on vacuolar phenolic compounds.20 Therefore, it can be seen that phenolic compounds and skin cell wall pectic fraction skeletons were little affected by the PEF1 treatment, and it can be confirmed that the polyphenols affected by high-intensity short duration PEF treatment are the vacuolar tannins rather than the cell wall tannins.20 In this way, PEF1 can be considered as a treatment that has little immediate destructive effect on cells, this effect remaining reversible, which explains the increase in skin size, without cell destruction. However, in the longer term, the skins of treated berries release their condensed tannins less easily, as they are strongly bonded to the affected pectic cell wall fractions. Their liberation takes place with a “delayed action effect”, which implies a lower rate of tannin extraction from the skins during maceration20 With regard to the PEF2 treatment, the skin structure of the treated berries was very largely degraded following the treatment. It was observed that the histological (Figure 1) and cytological (Figure 3) impacts on the cells were very significant. PEF2 was of lesser intensity, but was of longer duration than PEF1, which explains the more destructive effects of the PEF2 treatment.4,5,7 As berries are composed of heterogeneous tissues,23,25 with cells of differing shapes and sizes, PEF propagation cannot be uniform,3,7,13 which partly explains this result. Moreover, the increase in treatment energy (W = 31 Wh/kg) applied caused a higher degree of cell lysis19 than was the case for PEF1 (W = 4 Wh/kg). In parallel, TEM observations revealed skin cells with severely affected membranes and organelles, with distension of the cell walls and rupture of the middle lamella (Figure 2). These effects can be compared with the structural changes induced by PEF treatment on bacteria10,54 or yeast,6,11 in which a much greater critical electric field strength is required to obtain this result. This is mainly related to their diameter, which is much smaller, and to their cell wall thickness, which is greater than that of grape skin cells.3,7,13 The cytological structure of the cell walls was very largely modified by the PEF2 treatment, which explains the modified aspect of the cell walls observed with TEM (Figure 2). This finding was confirmed by biochemical studies in which analysis of the polysaccharide fractions revealed that the entire pectic skeleton had been disrupted. The whole polysaccharide mesh was greatly weakened, only

Moreover, the presence of the cell walls is an important factor in the spread of PEF. Few studies have examined the influence of PEF on the walls. Furthermore, Janositz and Knorr3 have demonstrated, with tobacco cells and protoplasts, the protective effect of cell walls vis-a-vis PEF treatment, having observed that protoplast cell size increased slightly after reversible permeabilization by PEF treatment with few pulses. In the present study, PEF treatment has been shown for the first time to have an effect on several cellular compartments such as cell walls in addition to the membranes: the skin of the treated berries was, overall, only slightly affected by the PEF1 treatment (4 kV/cm, 1 ms, 4 Wh/kg), but very largely degraded following the PEF 2 treatment (0.7 kV/cm, 200 ms, 31 Wh/kg). The most striking histological effect observed immediately after the PEF1 treatment concerned the thickness of the skin that was much greater than in the control batch, the most strongly affected cells being the large cells of the inner hypodermis.19,20 From a cytological point of view, skin-cell membranes of treated berries were found to be little affected by PEF1, which is logical in view of the PEF1 parameters (pulses of short duration (1 ms) and high strength (4 kV/cm)) that had induced electroporation on a lesser scale4 and which remained reversible. Conversely, cell walls of treated berries were more highly affected by PEF1 than membranes and, especially, the aspect of cell walls of the inner hypodermis immediately after the treatment (Figure 2). At the end of 14 days, cell walls were more distended in the inner hypodermis, showing certain areas with perforations in the outer hypodermis (Figure 3), which confirmed our findings that the consequences of PEF are not homogeneous.21−27 In this way, cytological observations revealed modifications of the polysaccharide skeleton interior, as did biochemical analysis of the pectic fractions (galacturonic acid, arabinose, and galactose).43 Pectic cell wall organization was only slightly but measurably affected further to the treatment: external fractions (PSE, PSOX) tended to increase, which probably increased the number of weak bonds,43 and the quantity of internal fractions (PSH, PSOH) was decreased, which also decreased the number of strong bonds. 43 It can be hypothesized that the disappearance of PSH and PSOH had disturbed the microfibrillar interactions of cell walls,29 thus freeing new space at the heart of the cell walls into which water may penetrate, thus inducing a stretching thereof. The distension of the walls that was observed immediately after the treatment would then allow the expansion of these cells, which would explain the thickening of the treated skin. The largest cells being the most strongly affected,3,7,13 this expansion was most noticeable at the inner hypodermis, which is made up of very large cells.23,25 Moreover, this upheaval of pectic skeleton has probably induced a change of affinity for the various parietal molecules such as polyphenols.47−50,53 With regard to the polyphenolic organization of the treated skin, biochemical analysis of the cell walls of total tannins in the different pectic fractions did not reveal any significant effect. However, a trend was observed: levels of TPSOX (inside the external pectic fraction) and TPSH (inside the internal pectic fraction) decreased, and their mDP increased. Following PEF1 treatment, PSOX and PSH fractions were depleted of total tannins because of the condensation of these tannins (mDP increased). It was possible to confirm this observation with TEM observations, in which structures resembling small balls were observed in the middle lamella walls (Figure 2). These structures may correspond to the condensation of TPSH in the space freed by the disappearance 2931



Article

facilitated polyphenol extraction. Thus, the choice of treatment modality is of prime importance because this choice can modify the kinetics of polyphenol extraction and, thus, wine quality. Moreover, it appears that there is a correlation between the mDP of tannins, astringency, and structure of wine, which has an influence on wine quality (low mDP tannins seem less astringent than strong mDP tannins).34 The availability of a technique that allows the extraction of more and better skin tannins during maceration or a technique that reduces maceration time, especially in the case of unripe harvests, and which, in addition, lead to a reduction of mDP (which is relatively high when the harvest is unripe), may be interesting from a qualitative point of view.34,38 This study is a first approach to the study of the impact of PEF on cell walls, and further research is required to better understand exactly which cell wall compounds are affected by PEF treatments and to better comprehend the mechanisms involved in improving the extractability of tannins by PEF treatment.

one internal fraction being strengthened. Except for PSOX, which increased, the quantities of all fractions were significantly decreased, which probably caused a loss of PSE, which is the most easily extractable fraction,42 and a reduction in the number of binding sites between polysaccharides and phenolic compounds.47,53 In the same way, biochemical analysis of total cell wall tannins demonstrated that the treatment profoundly modified the organization of the tannic skeleton. In the external fractions of the cell walls, TPSE were ruptured (mDP was decreased) contributing to the increase of their numbers in this fraction and probably inducing a decrease in their affinity for the polysaccharides of this fraction.53 At the same time, TPSOX became even more condensed (mDP was increased), which increased their affinity for the PSOX polysaccharide fraction, for which the numbers of weak bonds were increased. In the internal fractions of the cell walls, TPSOH were ruptured (mDP decreased) and remained in this fraction, whereas TPSH were also ruptured (mDP decreased) but migrated to the PSOH fraction in view of TPSH values, which decreased (Figure 2). Despite this enrichment in tannins, cell walls may have been weakened by a loss of the protective effect of the phenolic compounds against pectin depolymerization.32,53 The PEF2 treatment induced, on the one hand, an overall decrease in the number of weak bonds due to the loss of peripheral pectic fractions and loss of the strong bonds due to the loss of the internal pectic fraction and, on the other hand, a liberation of the majority of the cell wall tannins, which implies an overall decrease in the affinity of polyphenols for the cell wall fraction. In this way, the biochemical and physical structures of the cell walls were weakened by treatment, favoring the early release of cellular polyphenols, which explains why, at day 14, the skin vacuolar tannins of treated berries were depleted. The destructuring of the cell walls by the treatment facilitated the extraction and diffusion of phenolic compounds. This hypothesis seems to confirm the results of Delsart et al.19,20 concerning musts. These phenomena may have been facilitated at all layers of the skin by modification of the skeleton of tannins and the pectic fibers55,56 combined with ion leakage in cells and cell walls brought about by PEF treatment.7 Thus, low-intensity, long-duration PEF treatment profoundly modified the organization of the skin cell walls, which explains how the application of a PEF treatment enhanced phenolic compound extraction.15,18 This treatment mainly affected the cells wall tannins20 and severely affected cell wall protection. This suggests a more rapid release of cellular tannins due to the parietal system being disrupted and thus becoming more permeable to intracellular compounds. The extraction of polyphenols during maceration is facilitated by PEF2 treatment of the grapes. In conclusion, the comparison of the impact of two types of PEF treatment on the structure of grape berry skin allowed the very different consequences of the two treatment modes to be brought to light. In the case of PEF treatment of high intensity, short duration, and low energy, the effects on the structure of phenolic compounds and pectic cell walls of the skin were measurable but relatively limited. This study demonstrated a strengthening of the phenolic skeleton of skin cell walls, which has a protective effect, slowing polyphenol extraction phenomena. In the case of PEF treatment of higher strength, longer duration, and high energy, the consequences for the parietal structures of the skin were very significant, the pectic and phenolic skeletons being largely disorganized, which

■

AUTHOR INFORMATION

Corresponding Author

*(L.G.) Phone: +33 5 57 57 58 54. Fax: +33 5 57 57 58 13. Email: [email protected]. Funding

This work was supported by the Conseil Interprofessionnel des Vins de Bordeaux−33000 Bordeaux and by ADEME. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We thank Alan Metcalfe for corrections made to the text and Dr. Warren Albertin for expertise in statistical analysis.

■

ABBREVIATIONS USED eE, external epidermis; oH, outer hypodermis; iH, inner hypodermis; S, skin; VPC, vacuolar phenolic compounds; CW, cell wall; o, organelles; ml, middle lamella; p, plastid; m, mitochondria; mDP, mean degree of polymerization; PSE, extracting polysaccharide fraction in water; PSOX, extracting polysaccharide fraction in oxalate; PSH, extracting polysaccharide fraction in acid; PSOH, extracting polysaccharide fraction in sodium hydroxide; RP, residual-pellet polysaccharide fractions; TPSE, tannins of PSE; TSPOX, tannins of PSOX; TPSH, tannins of PSH; TPSOH, tannins of PSOH; TRP, tannins of RP; PEF1, treatment 1 of pulsed electric field; PEF2, treatment 2 of pulsed electric field; C, control berries; TEM, transmission electron microscopy; MO, optical microscopy

■

REFERENCES

(1) Chang, D. C. Cell poration and cell fusion using an oscillating electric field. Biophys. J. 1989, 56, 641−652. (2) Fincan, M.; Dejmek, P. In situ visualization of the effect of a pulsed electric field on plant tissue. J. Food Eng. 2002, 55 (3), 223− 230. (3) Janositz, A.; Knorr, D. Microscopic visualization of pulsed electric field induced changes on plant cellular level. Innovative Food Sci. Emerging Technol. 2010, 11 (4), 592−597. (4) Kandušer, M.; Šentjurc, M.; Miklavčič, D. The temperature effect during pulse application on cell membrane fluidity and permeabilization. Bioelectrochemistry 2008, 74 (1), 52−57. (5) Kandušer, M.; Miklavčič, D. Electroporation in biological cell and tissue: an overview. In Electrotechnologies for Extraction from Food

2932



Article

Plants and Biomaterials; Vorobiev, Lebovka, Eds.; Springer Science +Business Media: Dordrecht, The Netherlands, 2008; pp 1−37. (6) Harrison, S. L.; Barbosa-Cánovas, G. V.; Swanson, B. G. Saccharomyces cerevisiae structural changes induced by pulsed electric field treatment. Lebensm. Wiss. Technol. 1997, 30, 236−240. (7) Ersus, S.; Barrett, D. M. Determination of membrane integrity in onion tissues treated by pulsed electric fields: use of microscopic images and ion leakage measurements. Innovative Food Sci. Emerging Technol. 2010, 11 (4), 598−603. (8) Beveridge, J. R.; Macgregor, S. J.; Marsili, L.; Anderson, J. G.; Rowan, N. J.; Farish, O. Comparison of the effectiveness of biphase and monophase rectangular pulses for the inactivation of microorganisms using pulsed electric fields. IEEE Trans. Plasma Sci. 2002, 30, 1525−1531. (9) El Zakhem, H.; Lanoisellé, J. L.; Lebovka, N. I.; Nonus, M.; Vorobiev, E. Influence of temperature and surfactant on Escherichia coli inactivation in aqueous suspensions treated by moderate pulsed electric fields. Int. J. Food Microbiol. 2007, 120 (3), 259−265. (10) Chen, Y.; Yu, L. J.; Rupasinghe, H. V. Effect of thermal and nonthermal pasteurisation on the microbial inactivation and phenolic degradation in fruit juice: a mini-review. J. Sci. Food Agric. 2013, 93 (5), 981−986. (11) Huang, K.; Yu, L.; Liu, D.; Gai, L.; Wang, J. Modeling of yeast inactivation of PEF-treated chinese rice wine: effects of electric field intensity, treatment time and initial temperature. Food Res. Int. 2013, 54 (1), 456−467. (12) Angersbach, A.; Heinz, V.; Knorr, D. Effects of pulsed electric fields on cell membranes in real food systems. Innovative Food Sci. Emerging Technol. 2000, 1, 135−149. (13) Zimmermann, U. Electrical breakdown, electropermeabilization and electrofusion. Rev. Physiol., Biochem. Pharmacol. 1986, 105, 175− 256. (14) Grimi, N.; Lebovka, N. I.; Vorobiev, E.; Vaxelaire, J. Effect of a pulsed electric field treatment on expression behavior and juice quality on Chardonnay grape. Food Biophys. 2009, 4, 191−198. (15) Donsì, F.; Ferrari, G.; Fruilo, M.; Pataro, G. Pulsed electric fields − assisted vinification. Proc. Food Sci. 2011, 1, 780−785. (16) López, N.; Puértolas, E.; Condón, S.; Á lvarez, I.; Raso, J. Application of pulsed electric fields for improving the maceration process during vinification of red wine: influence of grape variety. Eur. Food Res. Technol. 2008, 227 (4), 1099−1107. (17) López, N.; Puértolas, E.; Condón, S.; Á lvarez, I.; Raso, J. Effects of pulsed electric fields on the extraction of phenolic compounds during the fermentation of must of Tempranillo grapes. Innovative Food Sci. Emerging Technol. 2008, 9 (4), 477−482. (18) Puértolas, E.; López, N.; Saldaña, G.; Á lvarez, I.; Raso, J. Evaluation of phenolic extraction during fermentation of red grapes treated by a continuous pulsed electric fields process at pilot-plant scale. J. Food Eng. 2010, 98 (1), 120−125. (19) Delsart, C.; Cholet, C.; Ghidossi, R.; Grimi, N.; Gontier, E.; Gény, L.; Vorobiev, E.; Mietton-Peuchot, M. Effects of pulsed electric fields on Cabernet Sauvignon grape berries and on the characteristics of wines. Food Bioprocess Technol. 2013, 1−13, DOI: 10.1007/s11947012-1039-7. (20) Delsart, C.; Ghidossi, R.; Poupot, C.; Cholet, C.; Grimi, N.; Vorobiev, E.; Milisic, V.; Mietton-Peuchot, M. Enhanced extraction of phenolic compounds from Merlot grapes by pulsed electric field treatment. Am. J. Enol. Vitic. 2012, 63 (2), 205−211. (21) Pratt, C. Reproductive anatomy in cultivated grapes − a review. Am. J. Enol. Vitic. 1971, 22 (2), 92−109. (22) Considine, J. A.; Knox, R. B. Development and histochemistry of the cells, cell walls and cuticle of the dermal system of the grape, Vitis vinifera L. Protoplasma 1979, 99, 347−365. (23) Hardie, W. J.; O’Brien, T. P.; Jaudzens, V. G. Morphology, anatomy and development of the pericarpe after anthesis in grape, Vitis vinifera L. Aust. J. Grape Wine Res. 1996, 2 (2), 97−142. (24) Ollat, N.; Diakou-Verdin, P.; Carde, J. P.; Barrieu, F.; Gaudillere, J. P.; Moing, A. Grape berry development: a review. J. Int. Sci. Vigne Vin 2002, 36 (3), 109−131.

(25) Fougère-Rifot, M.; Park, H. S.; Bouard, J. Ontogenèse du péricarpe de la baie de Vitis vinifera L. var. Merlot de la fécondation à la maturité. J. Int. Sci. Vigne Vin 1997, 31 (3), 109−118. (26) Fougère-Rifot, M.; Cholet, C.; Bouard, J. Evolution des parois des cellules de l’hypoderme de la baie de raisin lors de leur transformation en cellules de pulpe. J. Int. Sci. Vigne Vin 1996, 30 (2), 47−51. (27) Huang, X. H.; Huang, H. B.; Wang, H. C. Cell wall of loosening skin in post-veraison grape berries lose structural polysaccharides and calcium while accumulate structural proteins. Sci. Hortic. 2005, 104, 249−263. (28) Park, H. S.; Fougère-Rifot, M.; Bouard, J. Les tanins vacuolaires de la baie de Vitis vinifera L. var. Merlot à maturité. In Œnologie 95, 5ème Symposium International d’Œnologie; Tec&Doc: Paris, France, 1995; pp 115−118. (29) Goulao, L. F.; Oliveira, C. M. Cell wall modifications during fruit ripening: when a fruit is not the fruit. Trends Food Sci. Technol. 2008, 19, 4−25. (30) Pirie, A.; Mullin, M. G. Concentration of phenolics in the skin of grape berries during fruit development and ripening. Am. J. Enol. Vitic. 1980, 28 (4), 204−209. (31) Cholet, C.; Darné, G. Evolution of the contents in soluble phenolic compounds, in proanthocyanic tannins and in anthocyanins of shot grape berries of Vitis vinifera L. during their development. J. Int. Sci. Vigne Vin 2004, 38 (3), 171−180. (32) Gagné, S.; Saucier, C.; Gény, L. Composition and cellular localization of tannins in Cabernet Sauvignon skins during growth. J. Agric. Food Chem. 2006, 54 (25), 9465−9471. (33) Lacampagne, S. Localization and characterization of grape skin tannins: impact of cell wall physicochemical organization on tannin component, fruit quality and typicality of the Bordeaux grape. Ph.D. Thesis, Doctorat de l’Université Bordeaux2, 2010; 202 pp (http:// www.theses.fr/2010BOR21762/document). (34) Chira, K.; Pacella, N.; Jourdes, M.; Teissedre, P. L. Chemical and sensory evaluation of Bordeaux wines (Cabernet-Sauvignon and Merlot) and correlation with wine age. Food Chem. 2010, 126 (4), 1971−1977. (35) Colin, L.; Cholet, C.; Gény, L. Relationships between endogenous polyamines, cellular structure and arrested growth of grape berries. Aust. J. Grape Wine Res. 2002, 8 (2), 101−108. (36) McManus, J. F. A. Histological and histochemical uses of periodic acid. Stain Technol. 1948, 23 (3), 99−108. (37) Thiéry, J. P. Mise en évidence des polysaccharides sur coupes fines en microscopie électronique. J. Microscopie 1967, 6 (7), 987− 1018. (38) Reynolds, E. The use of lead citrate at high pH as an electron opaque stain in electron microscopy. J. Cell. Biol. 1963, 17, 208−212. (39) Gény, L.; Saucier, C.; Bracco, S.; Daviaud, F.; Glories, Y. Composition and cellular localization of tannins in grape seeds during maturation. J. Agric. Food Chem. 2003, 51 (27), 8051−8054. (40) Harris, P. J. Cell walls. In Isolation of Membranes and Organelles from Plant Cells; Hall, J. L., More, A. L., Eds.; Academic Press: New York, 1983; pp 25−83. (41) Deytieux-Belleau, C.; Vallet, A.; Donèche, B.; Geny, L. Pectin methylesterase and polygalacturonase in the developing grape skin. Plant Physiol. Biochem. 2008, 46 (7), 638−646. (42) Rapeanu, G.; Van Loey, A.; Smout, C.; Hendrickx, M. Thermal and high-pressure inactivation kinetics of polyphenol oxidase in Victoria grape must. J. Agric. Food Chem. 2005, 53 (8), 2988−2994. (43) Saulnier, L.; Thibault, J. F. Extraction and characterization of pectic substances from pulp of grape berries. Carbohydr. Polym. 1987, 7, 329−343. (44) Revilla, E.; Ryan, J. M.; Martin-Ortega, G. Comparison of several procedures used for the extraction of anthocyanins from red grapes. J. Agric. Food Chem. 1998, 46 (11), 4592−4597. (45) Kennedy, J. A.; Hayasaka, Y.; Vidal, S.; Waters, E. J.; Jones, G. P. Composition of grape skin proanthocyanidins at different stages of berry development. J. Agric. Food Chem. 2001, 49 (11), 5348−5355. 2933



Article

(46) Robertson, G. L. The fractional extraction and quantitative determination of pectic substance in grapes and musts. Am. J. Enol. Vitic. 1979, 30, 182−186. (47) Bindon, K. A.; Madani, S. H.; Pendleton, P.; Smith, P.; Kennedy, J. A. Factors affecting skin tannin extractability in ripening grapes. J. Agric. Food Chem. 2014, 62 (5), 1130−1141. (48) Hernández-Hierro, J. M.; Quijada-Morín, N.; MartínezLapuente, L.; Guadalupe, Z.; Ayestarán, B.; Rivas-Gonzalo, J. C.; Escribano-Bailón, M. T. Relationship between skin cell wall composition and anthocyanin extractability of Vitis vinifera L. cv. Tempranillo at different grape ripeness degree. Food Chem. 2014, 146, 41−47. (49) Ortega-Regules, A.; Romero-Cascales, I.; Ros-García, J. M.; López-Roca, J. M.; Gómez-Plaza, E. A first approach towards the relationship between grape skin cell-wall composition and anthocyanin extractability. Anal. Chim. Acta 2006, 563 (1−2 special issue), 26−32. (50) Renard, C. M. G. C.; Baron, A.; Guyot, S.; Drilleau, J. F. Interactions between apple cell walls and native apple polyphenols: quantification and some consequences. Int. J. Biol. Macromol. 2001, 29 (2), 115−125. (51) Saunder, J. A.; Lin, C. H.; Hou, B. H.; Cheng, J.; Tsengawa, N.; Lin, J. J.; Smith, C. R.; Macintoch, M. S.; Wert, S. V. Rapid optimization of electroporation conditions for plant cells protoplasts, and pollen. Mol. Biotechnol. 1995, 3, 181−190. (52) Wu, F. S.; Feng, T. Y. Delivery of plasmid DNA into intact plant cells by electroporation of plamolysed cells. Plant Cell Rep. 1999, 18, 381−386. (53) Le Bourvellec, C.; Guyot, S.; Renard, C. M. G. C. Non-covalent interaction between procyanidins and apple cell wall material: Part I. Effect of some environmental parameters. Biochim. Biophys. Acta−Gen. Subj. 2004, 1672 (3), 192−202. (54) Calderón-Miranda, M. L.; Barbosa-Cánovas, G. V.; Swanson, B. G. Inactivation of Listeria innocua in liquid whole egg by pulsed electric fields and nisin. Int. J. Food Microbiol. 1999, 51 (1), 7−17. (55) Barnavon, L.; Doco, T.; Terrier, N.; Ageorges, A.; Romieu, C.; Pellerin, P. Involvement of pectin methyl-esterase during the ripening of grape berries: cDNA isolation, transcript expression and changes in the degree of methyl-esterification of cell wall pectins. Phytochemistry 2001, 58, 693−701. (56) Cabanne, C.; Donèche, B. Changes in polygalacturonase activity and calcium content during ripening of grape berries. Am. J. Enol. Vitic. 2001, 5 (4), 331−335.

2934


Structural and Biochemical Changes Induced by Pulsed Electric Field

Recommend Documents