Relationship between antioxidant properties and chemical composition of the oil and the shell of pecan nuts [Caryaillinoinensis (Wangenh) C. Koch]

Industrial Crops and Products 45 (2013) 64–73

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Relationship between antioxidant properties and chemical composition of the oil and the shell of pecan nuts [Caryaillinoinensis (Wangenh) C. Koch] Ana Cristina Pinheiro do Prado a , Bruce A. Manion b , Koushik Seetharaman b , Francisco Carlos Deschamps c , Daniel Barrera Arellano d , Jane Mara Block a,∗ a

Federal University of Santa Catarina, Rod Admar Gonzaga, 1346, Itacorubi, Florianópolis, Santa Catarina, Brazil Department of Food Science – University of Guelph, Guelph, Ontario, Canada c Epagri Company of Agricultural Research and Rural Extension of Santa Catarina – Experimental Station of Itajai, Rod Anthony Heil, 06 km, Itaipava, Itajai, Santa Catarina, Brazil d State University of Campinas. University City Zeferino Vaz, Campinas, Sao Paulo, Brazil b

a r t i c l e

i n f o

Article history: Received 20 August 2012 Received in revised form 23 November 2012 Accepted 30 November 2012 Keywords: Pecan nut shell Chemical composition Antioxidant activity Infrared spectrometry Oxidation Oil Caryaillinoinensis

a b s t r a c t The chemical composition, oxidative stability, total phytosterols and tocopherols of pecan nut oil; the composition, color parameters and infrared spectroscopy of the pecan shell powder; the total phenolic compounds, condensed tannins and antioxidant activity (ABTS and DPPH) in extracts obtained by infusion of the pecan nut shell powder, were determined for samples of two consecutive years and different varieties. Furthermore, the possible effects caused by the use of spray drying technology were assessed. There was a significant effect for different harvest year and variety for sample quality and nutritional content of phytochemicals in the oil and nutshell. The results indicated a relationship between the content of unsaturated fatty acids present in the oil and the concentration of antioxidant compounds in the shell. Although phenolic compounds are considered to be thermally unstable, no significant losses were observed after the spray-drying process. © 2012 Elsevier B.V. All rights reserved.

1. Introduction The pecan nut [Caryaillinoinensis (Wangenh) C. Koch] belongs to the family Juglandaceae, originating from North America (USA). In the early twentieth century, its cultivation spread to several countries including Mexico, Australia, South Africa, Israel, Argentina and Brazil (Venkatachalam et al., 2007; Shahidi and Miraliakbari, 2005; Wakeling et al., 2001). In Brazil, the pecan nut was first introduced in the state of Sao Paulo. Currently Rio Grande do Sul is the largest producer of pecans in the country, followed by the states of Parana and Santa Catarina (Ortiz, 2000; De Carvalho, 1975). The basic nutritional composition of pecans produced in southern Brazil includes a high oil content (65–75%), and the predominant fatty acids include oleic (62.5%) and linoleic (27.5%).

∗ Corresponding author at: Federal University of Santa Catarina, Rod Admar Gonzaga, 1346 Itacorubi, Florianópolis, CEP: 88040-900, Santa Catarina, Brazil. Tel.: +55 48 3721 5367; fax: +55 48 3721 9943. E-mail addresses: [email protected] (A.C.P.d. Prado), [email protected] (B.A. Manion), [email protected] (K. Seetharaman), [email protected] (F.C. Deschamps), [email protected] (D. Barrera Arellano), [email protected] (J.M. Block). 0926-6690/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.indcrop.2012.11.042

Phytochemicals and other bioactive compounds, including tocopherols and phytosterols are also present in the pecan oil (Oro et al., 2008; Firestone, 1999). The tocopherols are aromatic compounds known for their vitamin E and antioxidant activity, which is strongly related to its chemical structure derived from the chromanol ring, and replaced by a grouping hydroxyl and methyl groups in the phenolic ring (Pokorny´ and Parkányiová, 2005; Wanasundara and Shahidi, 2005). The antioxidant activity of tocopherols is dependent on concentration, temperature, light, substrate and solvent, and lipid testing method, as well as the presence of synergists and chemical species that can act as pro-oxidants (Nogala-Kalucka et al., 2005; Bramley et al., 2000). Phytosterols are bioactive compounds that act on oxidative stability in plant cell membranes. They have in their chemical structure, a steroid nucleus with hydroxyl groups (3-␤-hydroxyl group), which may be related to the antioxidant activity exerted on the phospholipid bilayer of the cell membrane (Hounsome et al., 2008; Lagarda et al., 2006). In addition to the tocopherols and phytosterols in the pecan oil, other phenolic compounds such as phenolic acids and condensed tannins have been reported in pecan nut and nutshells using colorimetric assays (Folin-Ciocalteu and Vanillin) and of antioxidant activity assays (ABTS, DPPH, ORAC system and ␤-carotene

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linoleic acid) performed in vitro (Prado et al., 2009a, 2009b, 2010; Villarreal-Lozoya et al., 2007). In vivo studies also reported that phenolic compounds present in tea infusions from the shell of the pecan, can offer biological activity against drug-related diseases triggered by the action of free radicals preventing anxiety in withdrawal of the cigarette, and minimizing liver damage caused by oxidative stress in chronic alcohol ingestion (Müller et al., 2013; Reckziegel et al., 2011). The antioxidant activity of phenolic compounds occurs through the donation of a hydrogen atom from a hydroxyl group attached to the aromatic ring to the chemically unstable molecules, free radicals, thus slowing the oxidation process in cell membranes and foods rich in lipids. Moreover, they are capable of inhibiting the action of enzymes such as lipoxygenase, responsible for oxidative changes in fats and oils, improving the stability of such food products (Shahidi and Naczk, 2004). However, several factors may affect the antioxidant capacity of these compounds, including the processing conditions involving technology, solvent use and extraction temperature used in the isolation of phenolic substances (Moure et al., 2001). The concentration, the various chemical groups for the phenolic compounds, the species and variety of pecan being cultivated, in which antioxidants are synthesized naturally, can also significantly influence the characteristics such as shell color and astringent taste of tea obtained (Prado et al., 2010; Shahidi and Naczk, 2004). The pecan nut agricultural fields located in southern Brazil, comprised a mixture of varieties Barton, Shoshone, Shawnee, Choctaw and Cape Fear, and by 2008, it was estimated about 50% of the crops consisted of the Barton variety. This variety is obtained by grafting and breeding, and has received the most attention for large volume production, compared to other varieties of pecans due to its remarkable commercial importance (Divinut, 2011). In addition to variety, genetic factors, environmental conditions, soil composition, maturation and cultivation methods all have a strong influence on the phytochemical and nutritional composition of pecan nuts (Malick et al., 2009; Wakeling et al., 2001; Lavedrine et al., 2000; De Carvalho, 1975), and these factors may significantly affect the oxidative stability and antioxidant power of products from different regions and hemispheres. Although many studies have reported a simple relationship between the total phenolic content and antioxidant activity, this relationship can be much more complex in their contribution to the total antioxidant capacity, involving other matrix components in the plant, such as proteins, carbohydrates and fiber (Betancur-Ancona et al., 2004; Jing and Kitts, 2002; Amarovicz and Shahidi, 1997). Thus, in this study, the composition of the pecan nut oil and shell for two consecutive years (2009 and 2010) and different varieties (MV: a commercial mixture of varieties and B: Barton variety) were determined. The aim of this study, in addition to examining the effects of different harvest years and varieties on the composition of nuts, was also to evaluate a possible relationship between the general tenor of antioxidant compounds present in the oil and the shell of pecans, and the oxidative stability of the pecan nut oil. Furthermore, the possible effects caused by the use of spray dryer technology on the composition and color and antioxidant activity of powder extracts from the pecan shell were assessed.

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from Sigma–Aldrich (St. Louis, MO, USA); ABTS [2,2 -azino-bis(3-ethylbenzthiazoline-6-sulfonic acid)], DPPH (2,2-diphenyl-1picrylhydrazyl) and Trolox, obtained from Fluka/Sigma–Aldrich (St. Louis, MO, USA). All other chemicals and solvents used in the experiment were of analytical grade (PA) purchased from Vetec Fine Chemicals (Xerem, Rio de Janeiro, Brazil)/Sigma–Aldrich (St. Louis, MO, USA). 2.2. Samples of pecans Pecans harvested in the 2009 and 2010 growing seasons (6 kg for each harvest) from farms located in the central region of Rio Grande do Sul, Brazil were used. Samples studied consisted of a mixture of varieties including Barton, Shoshone, Shawnee, Choctaw, Stuart and Cape Fear, which is available commercially and designated as “MV”, and samples from crops consisting only of the variety Barton designated “B”. 2.3. Oil extraction of pecan nuts The nuts were shelled and the oil extracted by hydraulic press (model TE-098, Tecnal® , Sao Paulo, Brazil). The samples were pressed two times, and the oil was centrifuged at 3000 × g for 15 min. The supernatant was stored in amber bottles under nitrogen at −24 ◦ C until required for further analyses. The percentage extraction yield was calculated according to the equation: EY (%) = [(weight of extracted oil)/(weight of crushed almonds)] × 100, where EY = % extraction yield. 2.4. Obtaining the powder and extracts of the pecan shell The shells were dried at 40 ◦ C in an oven with forced air circulation (model 400/D 200 ◦ C, New Ethics® into reduce moisture content and then milled in a Mill analytical laboratory (model A-11 IKA Works® ). The powder was sieved to 60 mesh size and stored in amber bottles with nitrogen atmosphere at −24 ◦ C for later analysis (Prado et al., 2009a). The powder samples (MV and B) were placed in distilled water (20 g/L on a dry basis) and the extracts were stored according to procedure described by (Prado et al., 2009a). In addition, infusions were prepared according to Prado et al. (2009a, 2009b), using samples consisting only of the Barton variety of pecan shells (B) from the 2010 harvest. These extracts were dried by atomization spray drying using a BUCHI Mini Spray Dryer model B-290 (Buchi, Perdizes, Sao Paulo, Brazil). Spray drying conditions were as follows:air temperature inlet and outlet 150 ◦ C and 50 ◦ C, respectively, with aspirator set at 100% and pump at 25%) and stored at −24 ◦ C for later analysis (Sahin Nadeem et al., 2011). This sample powder was designated “B-SD”. The determination of the dry extract of all extracts was performed by gravimetric analysis, by pipeting an aliquot of 5 mL of extract into a pre-weighed and pre-dried aluminum pan, and drying in an oven (model 400/D 200 ◦ C, New Ethics® ) at 105.0 ± 0.5 ◦ C to constant weight (AOAC, 2005) to determine the extraction yield. 2.5. Chemical composition and oxidative stability of pecan oil

2. Materials and methods 2.1. Chemical reagents Standard fatty acid methyl esters (37 Sulpelco components Famex Mix, ref. U-47885) were obtained from Supelco – Sigma–Aldrich (Bellefonte, PA); Folin-Ciocalteau phenol reagent, gallic acid, vanillin, and (+)-catechin hydrated were obtained

2.5.1. Fatty acid composition Methyl esters were obtained according to the method of Hartman and Lago (1973) and the fatty acid composition was determined according to the methodology of AOCS Ce 1-91 (AOCS, 2004). Approximately 1 ␮L (split 1:50) of n-hexane phase was injected into gas chromatograph model Shimadzu GC-17A (Barra Funda, Sao Paulo, Brazil), equipped with a Supelco SP2340 capillary column (60 m × 0.25 mm × 0.2 ␮m) and flame ionization detector (FID). The

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injector and detector temperatures were 260 ◦ C and 240 ◦ C, respectively, using hydrogen (0.67 mL/min) as carrier gas. After injection, the column temperature was maintained at 120 ◦ C for 5 min and ramped at 4 ◦ C/min to 240 ◦ C and maintained for 10 min (total run time = 45 min). Standards of fatty acid methyl esters (FAMEs mix containing 37 components from Supelco, ref. U-47885) were injected under the same conditions and their retention times determined. The composition was determined by the standardization of the areas expressed as a percentage.

2.5.2. Tocopherols The levels of tocopherols were determined according to AOCS official method – Ce 8-89(2004) employing high performance liquid chromatography (HPLC). A Perkin Elmer Series 200 HPLC (Perkin Elmer, Perdizes, Sao Paulo, Brazil) with the following analytical conditions: isocratic pump Perkin-Elmer Series 200, a fluorescence detector Perkin-Elmer Series 200th; wavelength – Excitation 290 nm Emission 330 nm; Analytical Column – Hibar RT 250 mm × 4 mm Li Chrosorb Si 60, 5 mm; mobile phase – hexane/isopropanol (99/1) at 1.0 mL/min flow rate.

2.5.3. Total phytosterols The total phytosterols were determined according to the colorimetric method based on the enzymatic reaction of beta-sitosterol adapted from Allain et al. (1974) as follow: the oil sample (∼50 mg) was weighed in a dried, pre-weighed glass tube and the sample was saponified at 80 ◦ C for one hour with 5 mL of 0.5 M KOH in methanol. The non-saponified fraction was separated by adding 1 mL of water and 2 mL of hexane. A 0.6 mL aliquot of the top hexane phase was transferred to a test tube under constant air flow and temperature of 45 ◦ C for solvent evaporation. The unsaponified material was dissolved in 40 mL of isopropanol, and after vortex mixing, 3 mL of the enzymaticreagent for cholesterol (PP -cholesterol – Gold Analisa) was added and the sample incubated at 35 ◦ C for 15 min. After incubation period, the color intensity was measured at 500 nm with a FEMTO 700S UV spectrophotometer. A calibration curve was prepared from a commercial standard of phytosterols (ICN Biomedicals Brand. USA Inc. Cat Num. 102886, Lot No. 8312F). Samples were repurified and the purity was confirmed by gas chromatography as described in previous analyses.

2.5.4. Oxidative stability index and peroxide value The oxidative stability (Oil Stability Index – OSI) of the pecan oil was determined according to the official methodology Cd 12b92 from AOCS (2004), using the Oxidative Stability Instrument (Omnion, Rockland, MA) at 110 ◦ C with an air flow of 9 L/h and a sample weight of 5 g. The determination of the peroxide value of pecan oil was performed according to the official methodology Cd 8-53 of the AOCS (2004).

2.6. Composition, color and spectrophotometric analysis of the pecan shell 2.6.1. Proximate composition The composition of the pecan nut shell samples ground to a powder of the MV and B samples (both harvests 2009/2010), and the samples dried by spray dryer atomization by (B-SD, harvest 2010) was determined according to official methodology: moisture (925.09 – AOAC, 2005), protein (920.87 – AOAC, 2005), total lipids (920.85 – AOAC, 2005), crude fiber (Ba 6a-05 – AOCS, 1996) and minerals represented by ash content (923.03 – AOAC, 2005); carbohydrates were estimated by difference.

2.6.2. Instrumental color analysis For the instrumental color analysis, a Minolta Chromo Meter CR 400 (Minolta, Osaka, Japan) colorimeter, coupled to the DP-100 processor with illuminant D65 and an angle of 10 degrees, was used. The CIELAB (Commission Internationale de l’Eclairage) evaluation system was employed, with the color scale L*, a* and b*, where L* is the luminosity (zero for black and white is 100), a* is the color variation from green to red [from −80 to 0 is green (−a), from 0 to +100 is red (+a)], and b* is the color variation from blue to yellow [from −100 to 0 is blue (−b) is from 0 to +70 yellow (+b)]. Chroma (C*) indicating color saturation, i.e. the proportion in which the color is mixed with white, black or gray and hue (H expressed in angles with 0◦ as red, yellow as 90◦ , 180◦ as green and blue as 270◦ , corresponding to +a, +b,−a, and −b, respectively) were also determined. 2.6.3. Infrared spectrophotometric analysis (FTIR) Soluble and insoluble fractions were collected from the samples of the pecan nut shell powders (B and MV for harvest 2010) by weighing 0.5 g of powder and pipetting 10 mL of deionized water into 50 mL capped centrifuge tubes. The B-SD (harvest 2010) only had a soluble fraction since the process of making an infusion followed by spray-drying rendered this sample fully soluble in water. Sample centrifuge tubes were then sonicated at 55 ◦ C for 1 h, and then centrifuged at 6000 × g for 10 min at 4 ◦ C. The supernatant (soluble fraction) was decanted into pre-weighed large glass Petri dishes and dried overnight at 50 ◦ C. Likewise, insoluble pellet fractions were dried overnight at 50 ◦ C in separate pre-weighed glass dishes. Yields of soluble and insoluble fractions were determined by weighing Petri dishes with samples after drying was complete. Spectral analysis in the visible range (400–800 nm) was performed for the soluble fractions of B, MV and the B-SD (harvest 2010) powders with 25 mg of powder re-dissolved into 10 mL of deionized water to make a stock solution (0.25 mg/mL) and dilutions were prepared (0.025, 0.05, 0.0625, 0.0833, and 0.125 mg/mL solutions). Spectra were collected using a Biochrom WPA S800 visible photodiode array spectrophotometer (Terra Universal, Fullerton, CA, USA) and analyzed using Grafico software for all dilutions, and a standard curve was generated at 420 nm allowing quantification of the amount of soluble fraction in solution (R2 = 0.9964). The average absorbances of three replicates were taken for the standard curve. ATR-FTIR spectra of pecan nut shell powders (B, MV and B-SD – harvest 2010) including the whole powder, the soluble and insoluble fractions were collected on an Avatar 370 FTIR spectrophotometer (Thermo Scientific – Nicolet, Ottawa, ON, Canada) using a Golden Gate diamond attenuated total reflectance (ATR) accessory with a deuteratedtriglycine sulfate (DTGS) in the 4000–400 cm−1 infrared spectral range at room temperature. Each spectrum collected 32 scans at 4 cm−1 resolution and the average taken. Spectral analysis was conducted using OMNIC v7.3 software. A minimum of three spectra with <10% standard deviation was used for qualitative estimates after area normalized. 2.7. Total phenolics, condensed tannins and antioxidant activity of extracts from the pecan nut shell 2.7.1. Determination of total phenolic compounds The total phenolic content was estimated using the FolinCiocalteu colorimetric method with some modifications (Prado et al., 2009a; Budini et al., 1980). Aliquots (0.1 mL) of appropriate dilutions of the extracts were added to 0.5 mL Folin-Ciocalteau reagent. The reaction was neutralized with saturated 1.5 mL of sodium carbonate (75 g/L) and the volume adjusted to 10 mL by adding deionized water. After incubation for 2 h at room

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temperature, the absorbance of the resulting blue solution was measured at 764 nm with a spectrophotometer (model Hitachi model UV spectrophotometer, U-1800 (Konica Minolta, Ramsey, NJ, USA)). Quantification was performed using a standard curve generated with dilutions of a gallic acid stock solution and the results were expressed as GAE g−1 mg of dry weight (mg gallic acid equivalents/g of defatted sample). 2.7.2. Determination of condensed tannin content Condensed tannin content was determined using the procedure described by Price et al. (1978), and adapted by Villarreal-Lozoya et al. (2007). Aliquots of 1 mL of appropriate dilutions of the extracts were collected and placed into two separate test tubes (one for sample and one for blank). Then, 5 mL of vanillin reagent (0.5 g reagent and 200 mL of 4% HCl in methanol) was added to each sample, and 4% HCl in methanol added to blank. The test tubes were kept in darkness for 20 min and absorbance was measured on a spectrophotometer (Hitachi model UV spectrophotometer, U-1800, Ramsey, NJ, USA) at 500 nm. The results were expressed as CE g−1 mg of dry weight (mg catechin equivalents/g of defatted sample). 2.7.3. Antioxidant activity – ABTS method The ABTS [2,2 -azino-bis (3-ethylbenzotiazoline-6-sulphonic acid)] assay was carried out according to Re et al. (1999) with some modifications. After preparing the radical ABTS (7 mM – 0.03836 g ABTS dissolved in 10 mL deionized water), a potassium persulfate solution (2.45 mM – 10 mL and 10 mL ABTS persulfate) was mixed, homogenized and stored in an amber flask in darkness for a minimum of 16 h was prepared. For the sample, an aliquot of 200 ␮L of the radical formed was pipetted and diluted in 10 mL absolute ethanol, analysis grade. Absorbance measurements by spectrophotometer at 734 nm in 10 mm cuvettes were made to verify the optical density was around 0.700 ± 0.05 for a baseline (model UV spectrophotometer Hitachi, U-1800). An aliquot of 980 ␮L of the diluted radical was pipetted and transferred to a 10 mm (1 mL) cuvette, while measuring absorbance (A754 = A0 ), then adding 20 ␮L of the sample immediately, and mixed by inverting with parafilm for a few seconds. Trolox was used as the standard (0.13209 g/500 mL H2 O). The calculation of the radical inhibition percentage was made using the following formula: % radical inhibition = (1 − Af /A0 ) × 100, where, “A0 ” is initial absorbance and “Af ” is the final absorbance The calculations were made for each concentration of the samples analyzed to generate a dose–response curve. The results were expressed as ␮mol TEAC/g of dry weight (␮mol Trolox equivalent antioxidant capacity/g of defatted sample). 2.7.4. Antioxidant activity – DPPH method The DPPH (2,2-diphenyl-1-picrylhydrazyl) assay was performed according to Brand-Williams et al. (1995), followed by modifications, as described by Prado et al. (2009a) and Mensor et al. (2001). After preparing the 0.1 mM solution of the DPPH radical (0.03943 g DPPH dissolved in 10 mL of 80% ethanol), an aliquot of 2.9 mL was pipetted into test tubes with 0.1 mL of extract. The samples were kept in darkness for 30 min, and then absorbance was measured with a spectrophotometer (Hitachi model UV spectrophotometer, U-1800) at 515 nm. Trolox was used for the standard (150 mg/L). The calculation of the radical inhibition percentage was made using the following formula: % radical inhibition = (1− Af /A0 ) × 100, where, “A0 ” is initial absorbance and “Af ” is the final absorbance. The calculations were made for each concentration of the samples analyzed to generate a dose–response. The results were expressed as the TEAC g−1 mg of dry weight (mg Trolox equivalent antioxidant capacity/g of defatted sample).

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2.8. Statistical analysis The statistical analysis of the data was performed using the SAS “for Windows” v6.11 and Statistica® v7.0 software. Results were submitted to analysis of variance (ANOVA), Tukey test (p < 0.05), simple linear regression and principal component analysis (PCA). All analyses were carried out in triplicate.

3. Results and discussion 3.1. Chemical composition and oxidative stability of pecan oil The average yield of extraction, obtained from the pressing process of pecan kernels to obtain oil averaged 51% for both types of samples (MV and B). Oro et al. (2008), using the same process, found lower average oil yields (45%) to those reported in this paper, which can be explained by the use of two-step pressing in this study compared to only a single pressing performed by Oro et al. (2008). Table A.1 shows the results obtained for the fatty acid composition, content of tocopherols, phytosterols and oxidative stability of pecan oil for samples of different harvest years (2009, 2010) and varieties (MV and B). The results for the composition in fatty acids in pecan oil reported by Oro et al. (2008), show levels similar to those found in this work with oleic acid (62.5%), linoleic (27.5%), palmitic (5.6%) and stearic (2.8%), from pecan nuts grown on cultivated land in southern Brazil composed of a mixture of varieties (Barton, Shoshone, Choctaw and Cape Fear). The same authors reported levels of ␥-tocopherol of 30 mg 100 g−1 , and oxidative stability (OSI) and peroxide value determinations of 9.8 h and 0.55 mEq kg−1 , respectively (Oro et al., 2008, 2009); however, these authors did not observe significant levels for ␣-tocopherol. The results from this study were also similar to those reported by Firestone (1999) for the fatty acid composition (90% unsaturated fatty acids) and total phytosterols (0.1–0.29%). However, the reported levels by Firestone (1999) were significantly higher for ␣tocopherol (5–37 mg 100 g−1 ), with lower values for ␥-tocopherol (2–12.5 mg 100 g−1 ) when compared to nuts grown in southern Brazil. Shahidi and Miraliakbari (2005) reported 55% oleic fatty acid, 33% linoleic acid, 7% palmitic acid, 2% stearic, 1.0 mg 100 g−1 of ␣-tocopherol, 17.6 mg 100 g−1 of ␥-tocopherol, and 0.073% total phytosterols for the oil from pecan nuts grown in North America. Wakeling et al. (2001) also reported for the oil from pecans grown in Australia, mean levels below the present work ofoleic acid (55.3%) and significantly higher levels for linoleic acid (33%), with similar levels of palmitic acid (6.6%), stearic acid (2.5%), and linoleic acid (1.7%). These authors evaluated different varieties and harvest years, and attributed the observed differences in fatty acid composition to the different cultivation sites and varieties studied, and factors such as soil and climate. Villarreal-Lozoya et al. (2007) and Malick et al. (2009), evaluating the fatty acid composition of oil from pecan nuts grown in North America, also found significant differences related to cultivar and growing conditions, organic or conventional, applied to different crops. Venkatachalam et al. (2007) reported levels above 90% for unsaturated fatty acids in the oil from pecan nuts grown in Texas. These authors observed contents of oleic acid and linoleic acid from 52.5 to 74.1% and from 17.7 to 37.5% respectively. These results were higher than those reported in nuts grown in Brazil indicating that the location of cultivation may significantly influence the fatty acid profile of the pecan nut. Statistical analysis of the results indicate that the year of harvest was significant (p < 0.05) for the content of saturated fatty acids (MV 2010 > MV 2009 and B 2010 > B 2009), monounsaturated (MV

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Table A.1 MVa (2009)

Determination Palmitic acid (C16:0) (%) Stearic acid (C18:0) (%) Oleic acid (C18:1n9c) (%) Linoleic acid (C18:2n6c) (%) Total saturated fatty acids (%) Total monounsaturated fatty acids (%) Total polyunsaturated fatty acids (%) ␣-Tocopherol (mg 100 g−1 ) ␥-Tocopherol (mg 100 g−1 ) Total tocopherols (mg 100 g−1 ) Total phytosterols (%) Oxidative stability index – OSI (h) Peroxide value – PV (mEq.kg−1 )

Aa

4.7 2.5Aa 74.9Aa 17.8Aa 7.2Ab 74.9Aa 17.8Aa 1.1Aa 38.1Aa 39.2Aa 0.22Aa 7.4Ba 0.91Aa

± ± ± ± ± ± ± ± ± ± ± ± ±

Bb (2009) 5.0Aa 2.4Ba 75.9A a 16.7Ab 7.4Ab 75.9Aa 16.7Ab 1.1Aa 33.4Aa 34.6Aa 0.21Aa 10.3Aa 0.38Ba

0.2 0.1 0.6 0.4 0.2 0.6 0.4 0.1 0.4 0.5 0.01 0.1 0.01

± ± ± ± ± ± ± ± ± ± ± ± ±

MV (2010)

B (2010)

5.4Aa ± 0.1 2.3Aa ± 0.1 69.6Ab ± 0.6 22.6Aa ± 0.6 7.7Aa ± 0.1 69.6Ab ± 0.6 22.6Aa ± 0.6 1.1Ba ± 0.1 23.8Bb ± 0.5 24.9Bb ± 0.5 0.19Ab ± 0.01 10.6Aa ± 1.3 0.82Aa ± 0.08

0.1 0.01 0.2 0.3 0.1 0.2 0.3 0.1 4.1 4.2 0.01 2.9 0.02

5.3Aa 2.3Aa 69.2Ab 23.1Aa 7.7Aa 69.2Ab 23.1Aa 1.3Aa 26.0Aa 27.3Aa 0.19Ab 11.2Aa 0.44Ba

± ± ± ± ± ± ± ± ± ± ± ± ±

0.2 0.2 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.1 0.01 0.7 0.03

Capital letters in the same line for equal samples of the same harvest and same lowercase letters in the same line for the same type of sample and different harvests, no significant differences (Tukey test, p < 0.05), mean ± standard deviation (n = 3). a MV = mixture of varieties. b B = Barton.

2009 > MV 2010 and B 2009 > B 2010), polyunsaturated (B 2010 > B 2009), ␥-tocopherol (MV 2009 > MV 2010) and total phytosterols (MV 2009 > MV 2010 and B 2009 > B 2010). The crop variety also showed a significant effect (p < 0.05) on some characteristics of the pecan oil, for example the Barton variety showed the lowest levels of stearic acid for the 2009 growing season. Significantly higher levels of ␥-tocopherol and ␣-tocopherol were also observed in the oil extracted from Barton pecans in the 2010 harvest. Varietal effect was also significant in evaluating the oxidative stability (peroxide value and OSI), as the oil from Barton pecans exhibited more oxidative stability than the mixture of varieties for both 2009 and 2010. Superior oxidative stability and smaller peroxide values were observed in oil from pecan nuts grown solely of the Barton variety.

The samples atomized by spray drying had higher carbohydrate content (95.01%) with significantly lower levels of moisture (2.77%), minerals (1.68%), protein (0.43%) and total lipids (0.07%). Crude fiber was not detected in the composition of ground powder atomized by spray dryer. This result may be explained by the fact the powder of these samples was obtained by atomization of the infusion, which contained the water-soluble material present in the initial sample of raw pecan shell. Prado et al. (2009a) evaluated the composition of mixtures of varieties of pecan shells and reported levels similar to those obtained in this work for the fiber content (48.6%) and protein (2.2%). However, the authors observed a significantly lower level of carbohydrates (29.6%) and significantly higher moisture (16.8%) and total lipids content (1.1%). De Carvalho (1975) and Wakeling et al. (2001), analyzing different varieties of pecans from Brazil and USA, respectively, reported significant differences in the nutritional composition of the pecan kernels. The observed differences in the pecan shell composition can be attributed, according to Singanusong et al. (2003), to the different crop varieties, cultural practices, harvesting times and geographical locations. Moreover, genetic factors, soil composition and maturation can also significantly influence the nutritional composition (Malick et al., 2009; Wakeling et al., 2001). Statistical analysis of the pecan shell composition revealed that harvest year was significant (p < 0.05) for crude fiber, moisture, protein (MV 2010 > MV 2009 and B 2010 > B 2009), carbohydrates (MV 2009 > MV 2010 and B 2009 > B 2010) and total lipids (B 2009 > B

3.2. Proximate composition, color and spectrophotometric analysis of the pecan shell powder Table A.2 shows the results obtained for the composition and the color analysis of the pecan shell powder produced by milling (MV and B) and by atomization of the infusion of milled powder by spray dryer (B-SD), for different harvest seasons and varieties. According to the compositional analysis of the raw pecan shell from MV and B sample varieties from 2009 and 2010 harvest seasons, the crude fiber content represented the largest fraction of the shell (44.77–49.77%), followed by the carbohydrate content (34.33–41.41%), moisture (7.95–11.83%), protein (2.21–2.84%), minerals (0.88–1.85%), then total lipids (0.31–0.91%). Table A.2 MVa (2009)

Determination −1

Component (g 100 g

Color parameter

)

Proteín Moisture Total lipids Total Minerals Crude fiber Carbohydrates L* C* a* b* H

Bb (2009) Ab

2.58 7.95Bb 0.42Ba 1.63Aa 46.11Ab 41.41Aa 49.16Ab 18.29Aa 10.63Ba 14.88Aa 54.46Aa

± ± ± ± ± ± ± ± ± ± ±

MV (2010) 0.11 0.04 0.04 0.37 0.21 0.51 0.55 0.21 0.03 0.28 0.57

Bb

2.21 10.09Ab 0.91Aa 1.11Aa 44.77Bb 40.93Aa 47.43Ab 18.27Ab 11.79Aa 13.95Bb 49.79Bb

± ± ± ± ± ± ± ± ± ± ±

0.12 0.07 0.10 0.24 0.21 0.21 0.63 0.08 0.23 0.08 0.71

B-SDc (2010)

B (2010) Aa

2.84 11.30Aa 0.39Aa 1.85Aa 48.47Ba 34.57Ab 51.05Aa 18.60Aa 10.67Ba 15.12Ba 54.79Aa

± ± ± ± ± ± ± ± ± ± ±

0.11 0.04 0.003 0.20 0.43 1.33 0.28 0.29 0.00 0.18 0.33

2.79Aa 11.83Aa 0.31Bb 0.88Ba 49.77Aa 34.33Ab 51.35Aa 19.59Aa 11.31Aa 16.00Aa 54.73Aa

± ± ± ± ± ± ± ± ± ± ±

0.03 0.20 0.006 0.16 0.28 0.47 0.31 0.18 0.15 0.11 0.18

0.43** ± 2.77** ± 0.07** ± 1.67** ± Nd 95.01** ± 64.71** ± 22.73** ± 12.15** ± 19.19** ± 57.67** ±

0.02 0.12 0.02 0.04 0.09 0.06 0.13 0.08 0.09 0.06

L* = luminosity, C* = chroma, a* =  green to red, b* =  blue to yellow, H = hue angle. Nd = no detected. Means with the same superscript uppercase letter in the same line for samples of the same harvest and, with the same superscript lowercase in the same line for the same type of sample and different harvest, are not significantly different (Tukey test, p < 0.05); overall mean values ± SD (n = 3). B-SD (2010) = sample atomized by spray dryer, compared only the samples B (2010), “**”indicates significant difference (Tukey test, p < 0.05). a Mixture variety. b Barton. c Barton subjected to the process of spray dryer.

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Fig. A.1. FTIR spectra of the samples from 2010 harvest. A: pecan shell powders from Barton (B), mixture of varieties (MV) and Barton atomized by spray dryer (B-SD), B: pecan shell powders from Barton (B) and its soluble and insoluble fractions; C: pecan shell powders from the mixture of varieties (MV) and its soluble and insoluble fractions, D: soluble fraction of the pecan shell powders from Barton (B), mixture variety (MV) and Barton atomized by spray dryer (B-SD).

2010). The Barton variety had the lowest concentrations of fiber and protein, and the highest averages for moisture and total lipids in the 2009 growing season, and showed higher levels of fiber in the 2010 harvest. The mixture of varieties (MV 2010) showed higher levels for total minerals and lipids. For the content of minerals and carbohydrates, no statistically significant differences were observed between the harvest year and between cultivars, respectively. According to CIELAB color values obtained for powders determined by instrumental analysis, the sample atomized by spray dryer (B-SD 2010) was observed to have higher L* values (greater luminosity/light intensity), higher a* and b* (tending toward a red color). The same trend was observed when examining the tone with the hue angles all closer to yellow and red, as well as more saturated color/chroma (C*). Through statistical analysis of results obtained in the instrumental color parameters of the raw pecan shell powder, there were significant effects (p < 0.05 – Table A.2) exercised by the harvest year and variety studied. Crops showed differences for the parameters of brightness (MV 2010 > MV 2009 and B 2010 > B 2009), for color saturation, and a higher b* (B 2010 > B 2009). The differences observed between varieties were significant for the variation of red (a*) (B 2009 > MV 2009 and B 2010 > MV 2010), and yellow (b*) (MV 2009 > B 2009 and B 2010 > MV 2010) and for tint or hue (MV 2009 > B 2009). In Fig. A.1, the FTIR spectra for the samples studied can be observed for the 2010 harvest. According to the results, the B and MV samples showed similar spectra and are in agreement with the FTIR spectra of pecan nut shell other reported by other researchers (Vaghetti et al., 2010; Klasson et al., 2009; Guo and Rockstraw, 2007). However, slight differences were observed in samples MV and B with the Barton sample showing larger peaks in

the 2800–3000 cm−1 wavenumber range ascribed to the CH stretch in aliphatic compounds and aldehyde groups (Klasson et al., 2009) and could be due to higher lipid content and some compounds of the fiber fraction from these samples since they decrease in the soluble fraction. In comparison with the B and MV, the B-SD spraydried sample showed dramatic differences in FTIR spectra. The large peak from 3000–3600 cm−1 ascribed to the OH stretch and often attributed to bound water in the sample is larger in the spray dried sample, while the peak at 1740 cm−1 ascribed to the C O stretch of carboxylic acids, esters, ketones and aldehydes is shown to disappear. The peak at 1600 cm−1 ascribed to the phenyl (C C) bonds of the aromatic rings and the various peaks from 1000 to 1400 cm−1 often ascribed to the C O bond characteristic of carbohydrates or phenolics are dramatically increased both in the spray-dried sample and the soluble fractions of B and MV. In addition, the aliphatic CH stretch peaks in the 2800–3000 cm−1 range have disappeared with the spray-dried sample, yet are only reduced in the soluble fractions of B and MV. It was considered that the differences observed in the FTIR spectra of the spray-dried sample (B-SD) were due to the more soluble nature of the spray-dried sample possibly reflecting aheterogeneous spray-drying procedure of the sample. Therefore, the soluble fractions were extracted by making an infusion of the whole pecan nut shell powders (B and MV) to be compared with that of the insoluble fraction remaining as sediment The soluble and insoluble fractions of the B (30.2% ± 2.4%, 67.3% ± 1.6%)and MV (33.2 ± 1.4% and 63.2% ± 2.6%) samples gave total yields of 97.6% and 96.4% for the Barton variety (B) and mixture of varieties (MV) samples, respectively, with MV showing a slightly larger soluble fraction. Fig. A.1 (B, C, and D) display the FTIR spectra for the B and MV samples including their soluble and insoluble fractions. It can be observed

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Fig. A.2. A: visible spectra of the soluble fractions from the pecan nut shell powders of the mixture of varieties (MV) at various concentrations; B: standard curves generated for the B and MV soluble fractions and B-SD pecan shell powders at various concentrations measured at absorbance 420 nm. *Harvest 2010.

the insoluble fractions more closely resembled the whole powders while the soluble fractions more closely resemble the spray-dried samples (B-SD) showing the same absence of the 1740 cm−1 peak for C O stretch for the carbonyl group with relative increases in the O H stretch from the 3000–3600 cm−1 and 1600 cm−1 peak for the C C stretch in aromatics. However, differences between the spraydried sample (B-SD) and the soluble fractions exist with the soluble fractions displaying a more aliphatic nature with remaining peaks in the 2800–3000 cm−1 range and in general more sharper highly resolved peaks indicative of the FTIR spectra typical of smaller MW molecules as seen in the more resolved peaks at 1153, 1113, 1051, 957 and 943 cm−1 . This seems to suggest greater polymer incorporation into the spray-dried sample (B-SD). In addition, the visible spectrum was evaluated for the soluble fractions of the B and MV samples and the B-D (harvest 2010) at various concentrations (Fig. A.2 -A and B). The three sample powders after being re-dissolved in water showed a similar orange-redbrown color in solution. Fig. A.2 -A represents this color, displaying the absorbance spectra of the MV sample in solution at various concentrations. This was very representative of the spectra obtained for the B, MV and B-SD samples at the same concentrations. The absorbance at 420 nm was selected to model a molar extinction coefficient for application in rapidly quantifying the amount of dissolved solids in solution, as this wavelength showed a near maximum absorbance offering higher sensitivity coupled with a relatively flat plateau for robustness in that wavelength range and excellent regression coefficients for all samples tested. Fig. A.2 B shows the standard curves generated at 420 nm using cuvettes with a 1 cm path length. The slopes for the three samples are quite similar suggesting a common extinction coefficient can be used across these varieties of pecan nut shell powders with an average slope of 5.31 ± 0.056 absorbance units per mg/mL of pecan nut shell powder with an intercept of −0.044 ± 0.01 and a regression coefficient of 0.9969 ± 0.0008. This indicates future work for quantifying amounts of solids dissolved in the pecan nut shell infusions can be reliably determined rapidly by spectrophotometer when diluted to concentration ranges between 0 and ∼0.25 mg/mL. 3.3. Total phenolics, condensed tannins and antioxidant activity of extracts from the pecan shell powder Table A.3 presents the results obtained for dry matter total solids content, total phenolics, condensed tannins and antioxidant activity of extracts (infusion and infusion atomized by spray dryer and re-suspended in water) of the pecan shell powders of different harvests and varieties. The fact that the sample atomized by spray dryer (B-SD 2010) had higher levels of total phenolics and condensed tannins compared to the original sample (B 2010), coupled with the trend of a

greater red coloration (higher a*), observed in this sample might be explained in terms of oxidation occurring during the spray drying process according to Senter and Forbus (1978). In these reactions, flavonoid compounds such as leucocyanidin and leucodelphinidin are oxidized to phlobaphenes (condensed tannins with red color), with oligomeric or polymeric structure derived from the flavonoid intermediate flavan-4-ols. Moreover, according to these authors, to a lesser extent, the color changes observed in the sample atomized by spray dryer can be related to the formation of cyanidin and delphinidin. Prado et al. (2010), analyzing the color of pecan shells from different batches, noted that samples with red tones also showed higher levels of condensed tannins, suggesting a relationship with a higher concentration of phlobaphenesand the red color of the shell extract. Moreover, the superior results observed for antioxidant activity in the sample atomized by spray dryer are directly related to higher amounts of phenolic compounds concentrated in these samples. After statistically evaluating the effects exerted by harvest year and variety in relation to the color of the shell powders and results for dry matter, total phenolics, condensed tannins and antioxidant activity (ABTS and DPPH) of the extracts, it was observed that the samples with the highest mean for luminosity (L*) also showed significant effects of year of harvest for the content of total phenolics. The more yellow tinted powders (greater b* values) showed the same effects on the harvest year for antioxidant activity (both DPPH and ABTS) and the year of harvest had a significant effect on the dry matter content (MV 2009 > MV 2010). The samples that showed significant differences based on variety, for the tendency toward yellow (higher b*) also exhibited the same trend in respect to the dry matter content and total phenolics. For this effect of tint or hue for the 2009 harvest, it was also observed to have higher content of condensed tannins and antioxidant activity (DPPH), establishing a possible relationship between samples with hue angles (H) closest to 90◦ (red to yellow) and the antioxidant activity (DPPH), the latter being related to the content of condensed tannins. Prado et al. (2009a), evaluating the infusion from nutshells of a mixture of commercial varieties of pecan nuts, reported means significantly different for the content of dry extract (23 g 100 g−1 ), total phenolic compounds (138 mg GAE g−1 ), tannins condensates (43 mg CE g−1 ), and antioxidant activity in ABTS (1404 ␮mol TEAC g−1 ) and DPPH (385 mg TEAC g−1 ) systems. The differences observed between the results of Prado et al. (2009a) and the present study, besides being related to growing conditions, soil and environmental factors among others, may also be related to different harvest years (2006 vs. 2009/2010), and varieties comprising a different mixture of pecan samples (Barton, Shoshone, Shawnee, Choctaw and Cape Fear). Previous research, evaluating the color of pecan nuts, reported significant effects across different

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Table A.3

Determination −1

TS(g 100 g ) TPC (mg GAE g−1 ) CT(mg CE g−1 ) ACABTS (␮molTEAC g−1 ) ACDPPH (mg TEAC g−1 )

MVa (2009) Aa

36.09 130.30Ab 49.51Aa 1467.90Aa 590.86Aa

± ± ± ± ±

Bb (2009) 0.41 3.45 0.62 19.81 18.03

Ba

32.39 94.04Bb 43.19Ba 1333.10Ab 346.57Bb

MV (2010) ± ± ± ± ±

0.13 2.66 0.44 14.82 5.64

Bb

26.61 145.41Ba 39.09Aa 1723.60Aa 561.67Aa

± ± ± ± ±

B–SDc (2010)

B (2010) 0.52 8.16 4.31 12.15 64.09

Aa

32.12 181.49Aa 36.94Aa 1809.01Aa 612.24Aa

± ± ± ± ±

0.43 6.97 3.20 27.18 26.73

46.36** ± 0.50 590.78** ± 4.41 48.70** ± 1.50 4124.83** ± 57.09 1210.97** ± 25.24

Dry extract total solids TS = g 100−1 (gravimetric assay), total phenolic TPC = mg GAE g−1 (gallic acid equivalents) in dry weight (Folin-Ciocalteu assay), condensed tannins CT = mg CE g−1 (catechin equivalents) of dry weight (vanillin assay), AC = antioxidant capacity – ␮mol TEAC g−1 (Trolox equivalent antioxidant activity) in dry weight (ABTS), antioxidant capacity AC = mg TEAC g−1 (Trolox equivalent) in dry weight (DPPH – TEAC). Capital letters in the same line for equal samples of the same harvest and same lowercase letters in the same line for the same type of sample and different harvest, no significant differences (Tukey test, p < 0.05), mean ± standard deviation (n = 3). B-SD (2010) = sample atomized by spray dryer, compared with only samples B (2010), where “**” indicates significant difference (Tukey test, p < 0.05) between samples. a Mixture of varieties. b Barton. c Barton subjected to the process of spray drying process.

cultivars (Grauke et al., 1998; Silva et al., 1995) and years of harvest (Grauke et al., 1998) for tone and hue (H) values. 3.4. Relationship between the antioxidant properties and chemical composition of oil and the pecan nut shells To better understand the trends and relationships amongst the many variables for the different samples of pecan oil and pecan nut shell powder, a principal component analysis (PCA) was applied to the results obtained for MV and B of the harvests of 2009 and 2010 (Fig. A.3). According to the results obtained for composition (fatty acids, tocopherols, and total phytosterols) and oxidative stability (OSI and PV) of the oil (Fig. A.3 – A) together accounting for 90.61% of the variability of the data in the first two dimensions with components 1 and 2 (CP1 and CP2 ) accounting for 75.03% and 15.58% of the total variance, respectively. CP1 showed a strong correlation between C18:0 (stearic acid), TF (total phytosterols), C18:1n9c (oleic acid), C18:2n6c (linoleic acid), ␥-tocopherol and C16:0 (palmitic acid). Graphically, it is possible to observe a strong positive correlation between C18:0, TF, ␥-tocopherol and C18:1n9c and an inverse relationship with the content of C16:0 and C18:2n6c. Within the first component, it was also observed a strong inverse relationship between the content of oleic and linoleic acids among the groups of pecan nuts studied. The results are in agreement with earlier ˜ (1998), who evaluated studies by Toro-Vazquez and Pérez-Briceno the oil of pecan nuts native to central Mexico and found an inverse

correlation was highly significant (r = 0.976, p < 0.0001) between the levels of oleic and linoleic acids. The PCA for the pecan oil also indicated a strong relationship between the ␣-tocopherol content and oxidative stability (OSI and PV) of the oil, and as the content of ␣-tocopherol increased, PV decreased, improving the oxidative stability index (OSI). It was also observed a different distribution among the groups of nuts studied according to their chemical composition, and trends emerged for the harvest year and variety studied, with the harvest year being more influential a factor for the amount of different fatty acids while the variety was more influential for oxidative stability (B > MV). Rudolph et al. (1992) reported a relationship between oxidative stability of pecan oil to decreased concentrations of linoleic acid in accelerated oxidation studies, and found the best stability results for the Barton variety among the different cultivars Apache, Caddo, Choctaw, Comanche, Mohawk, Shawnee, Sioux, and Wichita. This relationship between the increase in the linoleic acid content and a decrease in oxidative stability was not detected in this study and thus suggests additional factors such as other components present in the oil that may offer antioxidant effects such as specific isomers of tocopherols and phenolics compounds. As well, the variety of the nut may exert significant influence on the stability of the oil. In the PCA analysis including multiple variables of the composition, color parameters, phytochemicals and antioxidant activity measured for the pecan shell powder samples B and MV (Fig. A.3 – B) for both 2009 and 2010 harvest years, the principal components 1 and 2 (CP1 and CP2 ) explained 88.55% of the total variance,

Fig. A.3. Graphical representation of PCA performed for the studied samples of the harvests of 2009 and 2010. A: pecan oil to the mixture of varieties (MV) and variety Barton (B). B: shell powder pecan MV sample (mixture of varieties) and Barton (B).

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with PC1 and PC2 accounting for 66.22% and 22.33% of the variance, respectively. PC1 showed a positive correlation between antioxidant activity by ABTS and DPPH, total phenolics, fiber, protein and color parameters L*, b*, C* and H. A strong inverse correlation of these attributes was observed with the levels of lipids and carbohydrates, in the same component (PC1 ). For PC2 , a positive correlation between the mineral content, dry matter and condensed tannins was observed, while the levels of condensed tannins and minerals were negatively correlated with the moisture content and color parameter a*, respectively. The PCA of the pecan shell powder made it possible to observe a different distribution between groups of pecan samples analyzed according to their composition and color parameters. The influence of crop and variety studied was remarkable in samples B and MV (2010). These samples showed groups with more homogeneous characteristics with each other, with emphasis on the Barton variety (2010) that showed higher values in most parameters assessed, distinguishing itself significantly from that of samples B and MV (2009), which showed the lowest values for total phenolics and antioxidant activity (DPPH and ABTS). Through principal component analysis (PCA) and the results of instrumental color analysis, it was possible to clarify some interferences observed in colorimetric analysis for total phenolics and condensed tannins. The Folin-Ciocalteau reagent, in addition to reduction by phenolic compounds is also sensitive to the presence of certain proteins (Zaia et al., 1998) which can overestimate the level of the compounds of interest. The vanillin assay, widely used for determination of condensed tannins may also suffer interferences thus overestimating results. According to Schofield et al., 2001, vanillin condensed tannins assay seems to be reactive with subunits of polymers tannins. Nevertheless, its lack of specificity for condensed tannins. Moreover, vanillin also reacts in the presence of some groups of proteins phenolic nature, due to the instantaneous reduction of vanillin free by chemical forces (hydrogen bonding and hydrophobic interaction), which is accentuated in foods with lower fat content (Chobpattana et al., 2000, 2002). Such interference can be better understood with the aid of instrumental analysis of color through the CIELAB system where the a* > 0 indicates the direction of staining for the reds. It is known that the red color is strongly related to the content of tannins, especially for the group of phlobaphenes (Prado et al., 2010; Senter and Forbus, 1978). For example, it was observed in MV (2009) samples that protein levels were significantly higher compared to those obtained for the B (2009) samples yet also having significantly reduced levels for lipids, favoring conditions for the Folin-Ciocalteau and Vanillin assays to result in overestimated values for total phenolics and condensed tannins. This result can be verified by analyzing the color parameter a* that reported values for redder hues in the B (2009) sample and principal component analysis in which relationships between the protein and lipid (strongly negative) and the parameter a* and the lipid content of the samples (strongly positive). The same magnitude of effect was not observed in the analysis of the other samples where the differences between the content of tannins was not significant. In samples atomized in spray dryer, there was a higher concentration of condensed tannin content, and a dramatic reduction in lipid content causing greater exposure of protein molecules, although these samples present a very low protein concentration. Furthermore, spray-dryed samples were more concentrated in phenolics. According to Shahidi and Naczk (2004), certain phenolic compounds present in the shell can be leached into the seed during the preconditioning by immersion of nuts into 80 ◦ C water before being broken during the stripping process. Senter et al. (1980) reported the quantity and quality of phenolic compounds are directly related to the oxidative stability of pecans during storage. This supports

the observation that although the 2010 harvest samples exhibited higher levels of polyunsaturated fatty acids, chemically more susceptible to oxidation reactions, they also showed higher values for oxidative stability of the oil. This suggests the possibility that pecan nuts with fatty acids more susceptible to oxidation may show higher levels of antioxidant compounds in their shells. Considering the link between both parts of the nut through the inner integument, which connects the shell to the kernel, the integument thereby seems to exert a protective effect in relation to oil in the kernel, due to preferential oxidation of phenolic compounds present in the shell (Jurd, 1956). This process would occur as a natural response to protect the plant to oxidative stress made more evident due to its composition of their nature more unsaturated fatty acids having higher measured oxidative stability and antioxidant activity. 4. Conclusions In this research, a significant effect for harvest year and variety of pecan sample was observed for the phytochemical and nutritional quality of the oil and nut shells. In addition, pecans harvested in 2010, in which higher levels were detected for polyunsaturated fatty acids (linoleic acid), also showed higher levels of total phenolics and antioxidant activity (DPPH and ABTS) in the shell as compared to the harvest of 2009, for same sample group. This suggests that the quantity and quality of phenolic compounds may bedirectly related to oxidative stability of the pecan oil, as evidenced by an observed relationship between the content of unsaturated fatty acids present in the oil and the concentration of antioxidant compounds in the protective outer shell. Although phenolic compounds are considered to be thermally unstable, there were no significant decreases in the spectrophotometric analysis in the infrared region (FTIR), the total phenolic concentration (CT and TPC), and antioxidant activity (DPPH and ABTS) after spray drying to a soluble powder offering a potential processing technique to concentrate these desired ingredients. Acknowledgments Divinut Walnut Ind. Ltd. (Cachoeira do Su, RS, Brazil) for providing the raw material, and Gum Products International (Newmarket, Ontario, Canada) for the use of the FTIR are gratefully acknowledged. Also, the authors would like to thank the Coordination for the Improvement of Higher Education Personnel (CAPES) for providing the scholarship for financial support, and Rafael Luchtenberg, Paula Cristina Engler, Priscilla de BritoPolicarpi, Priscilla de Oliveira dos Santos and Vanessa Martins Hissanaga, from Laboratory of Oils and Fats (UFSC), for their assistance in the analyses. This study was financially supported by CNPq process 479069/2011-5. References Allain, C.C., Poon, L.S., Chan, C.S.G., Richmond, W., Fu, P.C., 1974. Enzymatic determination of total serum cholesterol. Clin. Chem. 20 (4), 470–475. Amarovicz, R., Shahidi, F., 1997. Antioxidant activity of peptide fractions of capelin protein hydrolysates. Food Chem. 58, 355–359. AOAC, 2005. Official methods of analysis of the AOAC. In: AOAC – Association of Official Analytical Chemists, 18th ed. AOAC, Arlington, VA. AOCS, 1996. Approved method Ba 6a-05: crude fiber analysis in feeds by filter bag technique. Official methods and recommended practices. In: AOCS – American Oil Chemists Society, 4th ed. AOCS, Champaign, IL. 2004. Official methods and recommended practices of the American Oil Chemists Society. In: AOCS – American Oil Chemists Society, 5th ed. AOCS, Champaign, IL. ˜ Y., Fuertes-Blanco, S., Betancur-Ancona, D., Peraza-Mercado, G., Moguel-Ordonez, 2004. Physicochemical characterization of lima bean (Phaseoluslunatus) and Jack bean (Canavaliaensiformis) fibrous residues. Food Chem. 84, 287–295. Bramley, P.M., Elmadfa, I., Kafatos, A., Kelly, F.J., Manios, Y., Roxborough, H.E., Schuch, W., Sheehy, P.J.A., Wagner, K-H., 2000. Rewiew vitamin E. J. Sci. Food Agric. 80, 913–938.

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