Food Chemistry 136 (2013) 1277–1287

Contents lists available at SciVerse ScienceDirect

Food Chemistry

journal homepage: www.elsevier .com/locate / foodchem

Recovery and characterisation of coloured phenolic preparations from apple seeds

Matthias Fromm, Helene M. Loos, Sandra Bayha, Reinhold Carle, Dietmar R. Kammerer ⇑Hohenheim University, Institute of Food Science and Biotechnology, Chair Plant Foodstuff Technology, Garbenstrasse 25, D-70599 Stuttgart, Germany

a r t i c l e i n f o a b s t r a c t

Article history:Received 21 June 2012Received in revised form 6 September 2012Accepted 11 September 2012Available online 18 September 2012

Keywords:Apple seedsPhenolic compoundsExtractionOxidationPolyphenol oxidaseColourants

0308-8146/$ - see front matter � 2012 Elsevier Ltd. Ahttp://dx.doi.org/10.1016/j.foodchem.2012.09.042

⇑ Corresponding author. Tel.: +49 (0) 711 459 22995E-mail addresses: Dietmar.Kammerer@uni-hoh

hohenheim.de (D.R. Kammerer).

The aim of this study was to investigate whether complexly constituted phenolic extracts from appleseeds may be utilised for the recovery of natural coloured antioxidant preparations, which might serveas potential food or cosmetic ingredients.

In a first step, the recovery of phenolic compounds was optimised by varying crucial extraction para-meters. A single extraction step at 25 �C using an acetone–water mixture (60:40, v/v) and a solid-to-solvent ratio of 1:8 (w/v) for 1 h was found to be appropriate to achieve both high phenolic yields andantioxidant activities.

In a second step, differently produced apple seed extracts and a phloridzin model solution were enzy-matically treated by mushroom polyphenol oxidase to investigate the rate of pigment synthesis. Depend-ing on the extraction procedure applied, synthesis rates, pigment yields and colour propertiessignificantly differed. Compared to the phloridzin model solution, extracts recovered from the seedsshowed comparable and even better results, thus indicating such preparations to be a promising alterna-tive to synthetic yellow dyes.

� 2012 Elsevier Ltd. All rights reserved.

1. Introduction

Apples (Malus domestica Borkh.) belong to the world’s mostimportant fruit crops. In 2009, their global production accountedfor about 70 million tonnes (FAOSTAT, 2010). Approximately 70%thereof is marketed as fresh fruit, while 25–30% of the fruits areprocessed into juices or juice concentrates, ciders, purées, jamsand dried products. With an estimated production of 1.4 milliontonnes worldwide during 2004–2005, apple juice and concentratesthereof are the most important processed products (Bhushan, Kalia,Sharma, Singh, & Ahuja, 2008). Consequently, large quantities of ap-ple pomace are accumulated each year in the course of apple juiceproduction as agro-industrial by-products. So far, apple pomace hasbeen primarily used as animal feed (Vendruscolo, Albuquerque, &Streit, 2008) and for the recovery of pectins (Hwang, Kim, & Kim,1998). During recent decades, however, the recovery of phenoliccompounds from apple pomace is increasingly gaining importancedue to their antioxidant properties (Vanzani, Rossetto, De Marco,Rigo, & Scarpa, 2011; Vieira et al., 2011) and their potential healthbenefits (Bellion et al., 2010; Veeriah et al., 2006). While this devel-opment has led to numerous studies on phenolic compounds fromapple pomace (Cao, Wang, Pei, & Sun, 2009; Lu & Foo, 1997; Wijng-aard & Brunton, 2010), data on isolated apple seeds are ratherscarce or outdated. The studies reported so far have shown the phe-

ll rights reserved.

; fax: +49 (0) 711 459 24110.enheim.de, dietmark@uni-

nolic profile of apple seeds to be predominated by phloretin 20-xyloglucoside, 5-caffeoylquinic acid (chlorogenic acid), p-couma-roylquinic acid, (�)-epicatechin, some quercetin glycosides andphloridzin in particular. The latter was found to be the majorlow-molecular-weight phenolic constituent amounting to 60–90%of apple seed polyphenols (Fromm, Bayha, Carle, & Kammerer,2012a; Rupasinghe & Kean, 2008; Schieber et al., 2003). In severalstudies of recent years, phloridzin has been demonstrated to effec-tively inhibit the intestinal absorption and renal re-absorption ofglucose by the inhibition of sodium–glucose co-transporters 1and 2 (Alvarado & Crane, 1962; Chinard, Taylor, Nolan, & Enns,1957; Manzano & Williamson, 2010), thus providing a potentialmeans for the adjuvant treatment of diabetes mellitus type 2, obes-ity and stress hyperglycaemia (Ehrenkranz, Lewis, Kahn, & Roth,2005). Apart from its pharmacological properties, phloridzin andits aglycone phloretin have been identified as effective antioxidantsinhibiting lipid peroxidation (Rezk, Haenen, van der Vijgh, & Bast,2002; Rupasinghe & Yasmin, 2010).

Colour is one of the most important characteristics of foods andbeverages, strongly determining consumers’ acceptance. Sincesome of the synthetic azo dyes commonly applied in industrialfood processing have been associated with attention deficit hyper-activity disorder (ADHD) (McCann et al., 2007), they are nowincreasingly replaced by natural food colourants. In this context,an interesting characteristic of phloridzin has been reported uponoxidation by polyphenol oxidase (PPO). While phenolic compoundsof apple other than dihydrochalcones were oxidised to reddish-brown compounds, oxidation of phloridzin led to the formation

OGlc

HOCOOH

OHOH

OH

OGlc

HO OCOOH

O

OPOPi

POPj

Fig. 1. Structures of the phloridzin oxidation products POPi and POPj.

1278 M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287

of yellow products (Durkee & Poapst, 1965) which may be mainlyresponsible for the colour of apple juices and ciders (Lea, 1984).Hence, a range of studies on the formation and structures of thesewater-soluble phloridzin oxidation products (POP) have been con-ducted, and the formation of dimeric oxidation products was pro-posed (Goodenough, Kessell, Lea, & Loeffert, 1983; Oszmianski &Lee, 1991; Sarapuu & Kheinaru, 1969). However, only recentlythe structures of a colourless intermediate (POPi) and of a yellowpigment (POPj) (Fig. 1) could be elucidated by mass spectrometricand NMR spectroscopic experiments, revealing them to be mono-meric instead of dimeric oxidation products of phloridzin as postu-lated earlier (Le Guernévé, Sanoner, Drilleau, & Guyot, 2004). Insubsequent experiments the kinetics of their formation, their col-ouring properties and stability depending on pH value and temper-ature were investigated, indicating the potential use of POP asnatural food colourants (Guyot, Serrand, Le Quéré, Sanoner, &Renard, 2007).

Since the aforementioned studies were conducted with purechemicals in model solutions, the aim of the present study wasto investigate whether complexly constituted phenolic extracts de-rived from apple seeds are suitable for the recovery of colouredantioxidant preparations, which may serve as potential food orcosmetic ingredients. In a first step, the recovery of phenolic com-pounds should be optimised by investigating the effects of varyingextraction parameters on the contents and antioxidant activities ofapple seed extracts. In a second step, different extracts and a phlo-ridzin model solution should be enzymatically oxidised usingmushroom PPO to evaluate the efficiency of POPj pigmentsynthesis.

2. Material and methods

2.1. Chemicals

If not stated otherwise, all reagents and solvents were of analyt-ical or HPLC grade and were bought from VWR (Darmstadt, Ger-many). Ethanol (96%) was obtained from Nedalco Alcohol(Heilbronn, Germany). Phloridzin (purity > 95%) was obtained fromExtrasynthèse (Genay, France). 5-Caffeoylquinic acid was purchasedfrom Sigma–Aldrich (St. Louis, MO). Mushroom PPO (tyrosinase, EC1.14.18.1, 5370 U/mg), ABTS [2,20-azino-bis(3-ethylbenzothiazo-line-6-sulfonate)], Trolox [6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid], Folin reagent, ABAP [2,20-azobis(2-amidinopro-pane)dihydrochloride], TPTZ [2,4,6-tri(2-pyridyl)-s-triazine] andgallic acid were from Sigma–Aldrich (Steinheim, Germany).

2.2. Plant material

For most experiments, dried apple seeds kindly provided byHerbstreith & Fox (Neuenbürg, Germany) were used. They wererecovered from industrial dried apple pomace by sieving and man-ual separation from other pomace constituents. For some experi-ments freeze-dried apple seeds of different cider and dessertapple cultivars (harvest 2008) were used. Ripe apple fruits(‘Idared’, ‘Jonagold’, ‘Roter Ziegler’, ‘Boskoop’) were obtained from

Hohenheim University research station for horticulture (Stuttgart,Germany), local growers and from the local market.

2.3. Optimisation of phenolic compound extraction from apple seeds

2.3.1. Standard extraction protocolSeeds obtained from industrial apple pomace were ground con-

tinuously for 30 s using a laboratory mill and exhaustively ex-tracted with n-hexane to remove lipophilic constituents. Approx.2.5 g of the defatted apple seed flour were extracted for 2 h at25 �C using aqueous acetone (30:70, v/v; solid-to-solvent ra-tio = 1:8, w/v). To prevent oxidation of phenolic compounds, allextractions were carried out under nitrogen atmosphere. Sincepreliminary tests had shown stirring to be most effective, this pro-cedure was used throughout instead of shaking or sonication. Afterfiltration and washing of the residue, organic solvents were re-moved under reduced pressure (T < 30 �C). The recovered aqueousphenolic extracts were stored at �20 �C until further analyses.

2.3.2. Variation of extraction parametersTo optimise the extraction procedure for polyphenols from ap-

ple seeds, crucial parameters of the standard extraction protocol,such as type and proportions of solvents (ethanol, methanol, ace-tone) in water, solid-to-solvent ratios (1:8, 1:16, 1:24, w/v), extrac-tion time (1, 2, 3, 4, 8, 24 h) and temperature (0, 25, 42 �C) werevaried.

Moreover, the influence of different grinding techniques on thephenolic contents of the resulting apple seed extracts (ASEs) wasinvestigated. For this purpose, continuous grinding (‘30 s’) and suc-cessive grinding cycles with intermediate mixing (‘15 s + 15 s’;‘60 s + 30 s’) using a laboratory mill (model A 10, IKA, Staufen, Ger-many) were applied. For a better comparability the average parti-cle size distributions of the apple seed flours obtained weredetermined by sieve analysis. Additionally, a sieving mill (modelZM 1, Retsch, Haan, Germany) with a mesh size of 0.5 mm wasused for grinding (‘0.5 mm’). Polyphenolic extracts were obtainedafter an extraction time of 1 h applying the standard extractionprotocol.

Finally, the influence of the sample material itself was studiedby extracting apple seed flours from different authenticated applecultivars with aqueous acetone (30:70, v/v) for 1 h. Additionally,seed coats were manually separated from the seeds derived fromindustrial apple pomace. Both seed coats and de-hulled seeds (em-bryos) were separately analysed for their phenolic contents.

2.4. Enzymatic oxidation of phenolic compounds in extracts derivedfrom apple seeds

2.4.1. Recovery of phenolic extractsDefatted seeds from industrial apple pomace were ground

(5 � 10 s) with the aforementioned laboratory mill and extractedwith the respective solvents for 1 h at ambient temperature undernitrogen atmosphere (solid-to-solvent ratio = 1:8, w/v). After filtra-tion and centrifugation, respectively, the resulting extracts wereevaporated to dryness (T < 30 �C), and the residues were re-dissolved in phosphate buffer (pH 6.5, 100 mM). To assure compa-rable phloridzin concentrations in each of the samples, the extractswere diluted appropriately with phosphate buffer according toHPLC results. For the recovery of extracts A, B and C, aqueousacetone (30:70, v/v) was used as solvent. Whereas extract A wasdirectly subjected to enzymatic oxidation after removal of theorganic solvent, extracts B and C were obtained by subsequentliquid–liquid extraction of the corresponding aqueous crudeextracts with ethyl acetate at pH 7 and 1.5, respectively. The ethylacetate fractions were dried with anhydrous sodium sulfate andfiltered. Extract D was recovered using aqueous ethanol (50:50,

M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287 1279

v/v), while pure water was used for the extraction of phenolic com-pounds to obtain extract E.

2.4.2. Enzymatic synthesis of phloridzin oxidation productsThe aforementioned ASEs and a phloridzin model solution, each

containing approx. 0.5 mM phloridzin, were incubated in phos-phate buffer (pH 6.5) using mushroom PPO (30 U/mL). Incubationexperiments were carried out at 30 �C with magnetic stirring(410 rpm) under oxygen exposure (air). For monitoring the forma-tion of POPj, aliquots of the incubation medium (2 mL) were takenand immediately heated for 2 min at 95 �C to stop enzymatic reac-tions. Subsequently, samples were cooled on ice, membrane-fil-tered (0.45 lm) and kept at �20 �C until further analysis. Inpreliminary tests the complete inactivation of PPO under the heat-ing conditions applied was assured.

2.4.3. Colour analysisColorimetric measurements were carried out using a Perkin-El-

mer Lambda 20 spectrophotometer (Perkin-Elmer, Rodgau-Jüges-heim, Germany) equipped with UV WinLab software (version2.85.04) and WinCol software (version 2.05). L⁄, a⁄, b⁄, chroma[C⁄ = ((a⁄)2 + (b⁄)2)0.5] and hue angle [h� = arctan (b⁄/a⁄)] valueswere calculated from spectra recorded across a range of 380–780 nm using illuminant D65 and a 10� observer angle.

2.5. Determination of phenolic compounds and phloridzin oxidationproducts (POPj, POPi) by HPLC-DAD-ESI-MSn

HPLC analyses were carried out using an Agilent HPLC series1100 chromatograph (Agilent, Waldbronn, Germany) equippedwith ChemStation software, a model G1322A degasser, a modelG1312A binary gradient pump, a model G1329/G1330A thermo-autosampler, a model G1316A column oven, and a modelG1315A diode array detector. Spectra were recorded from 200 to600 nm (peak width 0.2 min; data acquisition rate 1.25/s). Individ-ual compounds were characterised by their retention times, UV/Visand mass spectral data.

LC–MS analyses were performed using the same HPLC systemas described above connected on-line to a Bruker (Bremen, Ger-many) model Esquire 3000 + ion trap mass spectrometer fittedwith an electrospray ionisation (ESI) source. Nitrogen was usedas drying gas at a flow rate of 9 L/min and as nebuliser gas at apressure of 40 psi. The nebuliser temperature was set at 365 �C.Negative ion mass spectra were recorded over the m/z range of50–2800.

Individual polyphenolics and phloridzin oxidation products(POPj) were quantitated using external calibration with corre-sponding reference compounds. Where standard compounds wereabsent, quantitation was achieved using the calibration curves ofstructurally related constituents applying a molecular weight cor-rection factor (Chandra, Rana, & Li, 2001).

2.5.1. System I (phenolic compounds without enzymatic oxidation)Separation of phenolic compounds in ASEs was performed

according to Schieber, Keller, and Carle (2001) with slight modifi-cations using a Phenomenex (Torrance, CA) Synergi 4u Hydro-RP80Å column (150 mm � 3.0 mm i.d.) equipped with a Security-Guard AQ C18 guard column, 4 � 2.0 mm i.d. (Phenomenex,Aschaffenburg, Germany) operated at 25 �C. The mobile phase con-sisted of 2% (v/v) acetic acid in water (eluent A) and of 0.5% aceticacid in water and acetonitrile (50:50, v/v, eluent B). The gradientprogram was as follows: 10% B to 55% B (50 min), 55% B to 100%B (10 min), 100% B to 10% B (5 min), 10% B (5 min). The flow ratewas 0.4 mL/min. The injection volume for all samples was 4 lL.Detection wavelengths were 280 nm (dihydrochalcones,

flavan-3-ols), 320 nm (hydroxycinnamic acids) and 370 nm (flavo-nols), respectively.

2.5.2. System II (enzymatically treated phenolic extracts)Phloridzin oxidation products (POPj, POPi), phloridzin, phloretin

20-xyloglucoside and chlorogenic acid in enzymatically treatedASEs were separated using a Waters SunFire C18 column(250 mm � 4.6 mm i.d., 5 lm particle size; Waters, Wexford, Ire-land) equipped with a Phenomenex guard column SecurityGuardAQ C18, 4 � 2.0 mm i.d. (Phenomenex, Aschaffenburg, Germany)operated at 25 �C. The mobile phase consisted of 2% (v/v) aceticacid in water (eluent A) and of methanol (eluent B). The gradientprogram was as follows: 5% B to 14% B (5 min), 14% B to 20% B(35 min), 20% B to 100% B (21 min), 100% B (2 min), 100% B to 5%B (1 min), 5% B (6 min). The flow rate was 0.8 mL/min. The injec-tion volume for all samples was 10 lL. Detection wavelengthswere 280 nm (dihydrochalcones, POPi), 320 nm (hydroxycinnamicacids) and 400 nm (POPj), respectively.

2.6. Synthesis and preparative isolation of phloridzin oxidationproducts (POPj)

For the synthesis of POPj, a solution of phloridzin (2.6 mM) inphosphate buffer (pH 6.5; 100 mM, 225 mL) was incubated atambient temperature under oxygen exposure (air) with vigorousstirring in the presence of mushroom PPO (31,104 U) for 24 h. Sub-sequently, aliquots thereof (2 mL) were incubated in derivatisationtubes at 95 �C for 3 min using a water bath to stop enzymatic reac-tions. After cooling on ice the aliquots were membrane-filtered(0.45 lm) and subjected to preparative HPLC. Isolation of POPjwas carried out using a Phenomenex C18 Aqua column(250 mm � 21.2 mm i.d., particle size 5 lm, pore size 125 Å). TheHPLC system (Bischoff, Leonberg, Germany) consisted of an LC-CaDI 22–14 system controller, two 2250 solvent delivery HPLCcompact pump modules, and an SPD-10AVvp UV/Vis detector (Shi-madzu, Kyoto, Japan). Data were processed with McDAcq32 Con-trol software (version 2.0) (Bischoff). The mobile phase consistedof 2% (v/v) acetic acid in water (eluent A) and 0.5% acetic acid inwater and methanol (30:70, v/v, eluent B). Phloridzin oxidationproducts POPi and POPj were eluted isocratically with 50% eluentB at ambient temperature and a flow rate of 6.0 mL/min. Monitor-ing was performed at 280 nm. Phloridzin oxidation products wereidentified by HPLC-DAD-MSn experiments according to mass spec-tral data previously reported (Le Guernévé et al., 2004). Chromato-graphic purity of the isolated POPj was P94% (HPLC, 280 nm).

2.7. Determination of total phenolic contents and of antioxidantactivities

All photometric measurements were performed in duplicatewith a Biotek microplate spectrophotometer (Power Wave XS, Bio-tek Instruments, Bad Friedrichshall, Germany) equipped with Gen5software (Ver. 1.04.5) and 96-well microplate cuvettes.

2.7.1. Folin–Ciocalteu assayTotal phenolic contents (TPCs) of the ASEs were determined

applying the Folin–Ciocalteu assay according to Singleton, Orthofer,and Lamuela-Raventos (1999) with some modifications. Briefly,50 lL of sample were mixed with 60 lL of Folin–Ciocalteu reagent(diluted with water, 1:6, v/v). After incubation for 3 min, 80 lL ofsodium carbonate solution (7.5%, w/v in water) were added andthoroughly shaken. Absorption measurements at 720 nm were per-formed after 60 min of incubation in the dark. Total phenolics areexpressed as gallic acid equivalents (mg GAE/g defatted matter).

1280 M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287

2.7.2. TEAC assaySpectrophotometric determination of the antioxidant activity

was performed as previously described (Van den Berg, Haenen,van den Berg, & Bast, 1999). For the daily preparation of the ABTSradical reagent, 0.5 mL of ABTS solution (20 mM in phosphate buf-fer, pH 7.4) and 100 mL of ABAP solution (2.5 mM in phosphatebuffer, pH 7.4) were blended and heated for 15 min at 60 �C in awater bath in the dark. The phosphate buffer was prepared byblending 812 mL of a Na2HPO4�2H2O solution (66 mM) with182 mL of KH2PO4 solution (66 mM) and 8.8 g sodium chloride.Antioxidant activity was measured by mixing 40 lL of sample with200 lL of radical solution in microplate cuvettes. Absorption wasmeasured after 6 min at 734 nm.

2.7.3. FRAP assayAntioxidant activity was determined according to a method

previously reported (Benzie & Strain, 1996). FRAP reagent was pre-pared daily by blending 2.5 mL of TPTZ solution (10 mM in 40 mMHCl), 2.5 mL aqueous FeCl3 solution (20 mM) and 25 mL of an ace-tate buffer (300 mM, pH 3.6). Sample aliquots (20 lL) were mixedwith 150 lL of FRAP reagent, and absorption was measured after4 min at 593 nm.

Aqueous Trolox solutions were used for calibration of both theTEAC and FRAP assay. Antioxidant activities were calculated aslmol Trolox antioxidant equivalents (TAE)/g defatted matter.

2.8. Statistical analysis

All experiments were performed at least in duplicate. Signifi-cant differences (a = 0.05) were determined using the Tukey or Tu-key–Kramer test for different independent samples. Dataevaluation was performed with the SAS software package (Soft-ware Version 9.1; SAS Institute, Cary, NC).

3. Results and discussion

3.1. Optimisation of polyphenol extraction from apple seeds

The effects of varying crucial extraction parameters on the effi-ciency of polyphenol extraction from apple seed residues were as-sessed by means of Folin–Ciocalteu, TEAC and FRAP assays. Some ofthe extracts were additionally analysed by RP-HPLC.

3.1.1. Extraction solventsIn general, polyphenol recovery was strongly determined by the

solvent chosen for extraction of the plant material. Fig. 2a illus-trates the influence of varying acetone/water ratios on the totalphenolic contents (TPCs) of the extracts obtained. Maximum TPCsamounting to 2.99 and 2.89 mg GAE/g were found for 60% and 70%acetone, respectively, whereas the extraction with 100% acetoneproved to be most ineffective, since the respective extract was de-void of phenolics as determined by the Folin–Ciocalteu assay.When using pure water for extraction, TPC was also very low(0.96 mg GAE/g). These findings are in agreement with a previousstudy (Al-Farsi & Lee, 2008), indicating pure acetone and waterto be unsuitable extraction solvents. The TPC values (Fig. 2a)showed good correlation with the antioxidant activities summa-rised in Table S1. Maximum antioxidant activities were measuredin extracts obtained with an acetone–water mixture containing60% acetone (TEAC: 42.57 lmol TAE/g, FRAP: 6.42 lmol TAE/g).Antioxidant properties of the extracts obtained with pure acetonewere much lower (3.30 TAE/g lmol), displaying only about one-tenth of the maximum TEAC value, while antioxidant activitywas unverifiable using the FRAP assay. Additional HPLC analysis re-vealed insignificant difference with respect to phloridzin contents;

yielding 2.80 and 2.61 mg/g for the extractions with 60% and 70%acetone, respectively (Table S1). Since differences in the yields ofphenolic compounds and antioxidant activities were insignificantusing these two solvent compositions, both were considered tobe the most suitable extraction solvents for apple seed phenolics.

Regarding their TPCs, extraction efficiencies with differenthydroalcoholic solutions were very similar (Fig. 2b). Highestextraction yields were obtained both with 50% methanol(2.04 mg GAE/g) and 50% ethanol (2.01 mg GAE/g) in water. As ob-served for acetone, TPC of the extract recovered with pure metha-nol was significantly lower (0.91 mg GAE/g), reaching only half thevalue as compared to the extract obtained with the optimum sol-vent composition. Antioxidant activities showed a similar trend.Whereas TEAC and FRAP values were comparable for differenthydroalcoholic solvents, values for extracts obtained with puremethanol were significantly lower.

Comparison of all solvent systems evaluated in this study re-vealed acetone (60–70%, v/v) to be the optimum extraction med-ium for polyphenol recovery from apple seeds, both with regardto TPCs and antioxidant activities. This is consistent with the find-ings of a previous study demonstrating that aqueous acetone ex-tracts of apple seeds exhibit higher TPC values than thoseobtained by ethyl acetate or hydromethanolic extraction(González-Laredo et al., 2007). Phloridzin levels ranging from2.33 to 2.80 mg/g for the extracts obtained with 50–70% of organicsolvent, however, did not markedly differ. Thus, each of theseextraction mixtures may be similarly used for the efficient recov-ery of phloridzin.

3.1.2. Extraction time and solid-to-solvent ratioExtraction time and solid-to-solvent ratio are also known to

influence extraction yields. As illustrated by Fig. 2c the effect ofextraction time on TPCs of the ASEs was not significant. Whenextending extraction times up to 24 h, TCPs remained almost con-stant, varying between 3.00 (1 h) and 3.28 mg GAE/g (8 h). Similarobservations applied to the corresponding antioxidant activities,ranging from 5.54 to 7.29 lmol TAE/g and from 42.47 to46.53 lmol TAE/g for the FRAP and TEAC assays, respectively. Con-sequently, a minimum extraction time of 1 h was considered mostsuitable. In Fig. 2d TPCs of ASEs obtained using different solid-to-solvent ratios in combination with different extraction times arepresented. With decreasing solid-to-solvent ratios from 1:8 to1:24 TPCs were augmented for each of the extraction periods. Asshown for the extraction over 3 h, TPCs increased most signifi-cantly by 25% from 2.77 to 3.46 mg GAE/g. TPC gains were less pro-nounced, only amounting to 17% and 14%, when extraction timesof 1 and 2 h, respectively, were applied. Again, extraction time onlyslightly improved polyphenol extraction yields by 12% at maxi-mum from 2.96 to 3.32 mg GAE/g at a solid-to-solvent ratio of1:16 (w/v). Thus, in agreement with a previous report (Al-Farsi &Lee, 2008), decreasing the solid-to-solvent ratio proved to be morerelevant than extension of extraction time.

3.1.3. TemperatureTemperature may have great influence not only on the solubil-

ity of a solute, but also on its stability. In our experiments the effectof different temperatures on extraction yields was investigated.Since acetone–water was shown to be most efficient, this extrac-tion medium was used in subsequent experiments. Due to thelow boiling point of acetone (56 �C), temperature effects were eval-uated between 0 and 42 �C. As illustrated by the results for TPCs(Fig. 2e), extraction was most efficient at ambient temperature(25 �C), yielding a significantly higher TPC value (2.86 mg GAE/g)than for lower and higher temperatures (0 �C: 2.52 mg GAE/g;42 �C: 2.32 mg GAE/g). These findings correlated with the resultsof the TEAC and FRAP assays, since the extract obtained at 25 �C

(a) (b)

(c) (d)

(e)

(g)

(f)

Fig. 2. Effects of varying crucial extraction parameters (aqueous acetone (a), aqueous methanol or ethanol (b), extraction time (c), solid-to-solvent ratio (w/v) (d),temperature (e), comminution (f), sample material (g)) on total phenolic contents (mg GAE/g defatted matter) of extracts recovered from apple seeds. Bars represent means ofat least two determinations ± standard error. Identical letters indicate that values are not significantly different.

M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287 1281

exhibited highest antioxidant activities. Thus, low temperaturesobviously negatively affected the solubility and extraction of phen-olics, whereas elevated extraction temperature appeared to beassociated with progressive oxidative losses. Furthermore, due tosolvent evaporation at elevated temperatures, the acetone-to-water ratio may have changed which might have negatively af-fected extraction yields, as reported earlier (Durling et al., 2007).

3.1.4. Effect of sample comminutionSince extraction yields are strongly dependent on the particle

size of the sample material, and consequently on the comminutionprocedure applied, different grinding protocols, such as continuousgrinding (‘30 s’) and successive grinding cycles with intermediatemixing (‘15 s + 15 s’; ‘60 s + 30 s’) were studied using a laboratorymill. Alternatively, a sieving mill (‘0.5 mm’) was used. Fig. 2f sum-marises the TPCs of the resulting extracts. With values of 3.31 and

3.07 mg GAE/g, highest TPCs were determined when grinding ap-ple seeds for ‘60 s + 30 s’ and ‘30 s’, respectively. This is in agree-ment with their average particle size distribution showing highproportions of particles with minimum size <0.1 mm (6.6% and17.1%, respectively). Subsequent grinding for 15 s with intermedi-ate mixing (‘15 s + 15 s’) resulted in a significantly lower TPC of theextract (2.66 mg GAE/g), although 83.2% of the particles ranged be-tween 0.315 and 1.0 mm. Interestingly, particle sizes of the lattersample were not below 0.1 mm, thus confirming this particle sizeto be of utmost importance for efficient polyphenol extraction.Using a sieving mill, TPC of the corresponding extract amountedto 2.76 mg GAE/g. HPLC analysis revealed an appreciable phlorid-zin content of 2.96 mg/g, thus indicating the sieving mill to be analternative to the laboratory mill. However, it is worth mentioningthat during grinding with the sieving mill a marked raise of tem-perature was observed, which might have favoured oxidative

1282 M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287

polyphenol losses and aggregation of lipid-containing particles.Consequently, in order to optimise grinding of apple seeds, coolingis a prerequisite. For the extract obtained from apple seed flour‘60 s + 30 s’, antioxidant activities of 7.25 lmol TAE/g (FRAP) and46.47 lmol TAE/g (TEAC) were determined, thus exhibiting mark-edly higher antioxidant activity than the remaining samples, whichhad comparable antioxidant properties independent of the grind-ing procedure.

3.1.5. Sample materialSample material itself may also play an important role for

extraction yields. While there are several studies dealing withthe phenolic contents of apple fruits showing cultivar-dependentdifferences (Keller, Streker, Arnold, Schieber, & Carle, 2001; Tsao,Yang, Young, & Zhou, 2003), data on apple seeds are scarce. In or-der to fill this gap, seed flours of different apple cultivars, seedcoats and de-hulled seeds should be analysed for their phenoliccontents and antioxidant activities. As apparent from Fig. 2g, TPCsof the seeds of the different cultivars significantly differed. Highestcontents were determined for seeds of the cvs. ‘Boskoop’ and ‘RoterZiegler’, which contained 16.19 and 9.36 mg GAE/g, respectively. Incontrast, seeds of cv. ‘Jonagold’ exhibited a much lower TPC, onlyaccounting for 18% of the phenolic content of cv. ‘Boskoop’. Inaccordance with the respective pulps and peels, seeds of ciderapple cultivars contain higher levels of polyphenols than thoseof dessert apples. As observed for TPCs, phloridzin contents alsodiffered significantly among the seeds of different cultivars,ranging between 2.60 and 16.41 mg/g for the cvs. ‘Jonagold’ and‘Boskoop’.

Investigations into different apple seed tissues revealed theseed coats to exhibit a TPC of 8.10 mg GAE/g, whereas only0.20 mg GAE/g was found for the seeds. Taking into account theaverage proportion of the seed coats (�37%), coat phenolics ac-counted for 96% of TPC of the whole seeds. Consequently, seedpolyphenols are almost exclusively deposited in the seed coat tis-sue, while the fat-containing embryos are mostly devoid of suchpolar compounds. Since some of the cultivars, in particular ‘Bosko-op’, exhibited a significantly lower seed coat-to-embryo ratio, thedistribution of phenolics appears to be an additional reason forthe observed quantitative differences. These results were corrobo-rated by HPLC, revealing a negligible phloridzin content (0.10 mg/g) in the embryos. Interestingly, the phloridzin level of the seedcoat (4.92 mg/g) was approx. half the amount determined by theFolin–Ciocalteu assay. The observed discrepancy may be ascribedto the fact that monomeric, oligomeric and polymeric phenolicsare assessed by the photometric assay, while only low-molecular-weight phenolics, particularly phloridzin, are covered by RP-HPLC.Antioxidant activities of the respective ASEs showed good correla-tion with their TPCs, with cv. ‘Boskoop’ exhibiting the highest TEACand FRAP values, whereas cv. ‘Jonagold’ showed the opposite.

3.2. Identification of apple seed phenolics by HPLC-DAD-MSn

Identification of individual phenolic constituents was achievedby comparing their retention times, UV/Vis and mass spectral datawith those of reference compounds. Fig. 3 shows typical chromato-grams recorded at different wavelengths. Retention times, UV/Visand mass spectral data as well as peak assignments are specifiedin Table 1. At 280 nm a hydroxybenzoic acid, flavan-3-ols and mostof all compounds exhibiting dihydrochalcone structures were de-tected. Compound 1 revealed an [M�H]� at m/z 153 producing afragment ion at m/z 109 in MS2 experiments, thus indicating theloss of a carboxylic function. According to its UV/Vis and massspectra including its retention time, compound 1 was identifiedas protocatechuic acid. Among the flavan-3-ols, compounds 2and 3 were identified as procyanidin B2 and (�)-epicatechin,

respectively. In a ‘Boskoop’ seed extract, however, an additionalcompound (4) occurred, showing UV/Vis spectral data typical forprocyanidins and an [M�H]� ion at m/z 865. In MS2 and MS3 exper-iments fragment ions previously reported for a procyanidin trimerof the B-type were obtained (Verardo, Bonoli, Marconi, & Caboni,2008; Xiao et al., 2008). Reports on flavan-3-ols in apple seedsare quite contradictory. Whereas some authors identified (�)-epi-catechin and procyanidin B2 (Schieber et al., 2003) others detected(�)-epicatechin and (+)-catechin (Rupasinghe & Kean, 2008) oreven reported an absence of flavan-3-ols (Lu & Foo, 1998). Thesediscrepancies may be explained by different analytical methodsand differing raw materials.

According to their elution order and mass spectral behaviour,peaks 6, 7, 8 were assigned to the dihydrochalcones 3-hydroxy-phloridzin (m/z 451), phloretin 20-O-xyloglucoside (m/z 567) andphloridzin (m/z 435), respectively. Upon collision-induced dissoci-ation (CID), compounds 7 and 8 generated a fragment at m/z 273,indicating the loss of their saccharide moieties and the release ofthe phloretin aglycone. Compound 6, however, provided an ion atm/z 289, indicating the presence of an additional hydroxyl group(D = 16 Da). A similar fragmentation pattern has been previouslyreported in positive ionisation mode (Alonso-Salces et al., 2004).Eluting shortly before compound 6, compound 5 was detected,providing an [M�H]� ion at m/z 597. Its fragmentation pathwaywas almost identical to that of phloridzin. Furthermore, fragmentsat m/z 435 (MS2) and m/z 273 (MS3) were generated. The formationof these fragments was ascribed to two successive losses of a hex-ose moiety (162 Da each). Moreover, the UV/Vis spectrum of thiscompound showed maxima at 280 and 230 nm, thus being similarto a previously reported unknown dihydrochalcone or flavanonestructure (Lu & Foo, 1998). Based on these findings, compound 5was concluded to represent a phloretin dihexoside. Compound 9was identified as 5-caffeoylquinic acid by its UV/Vis spectral dataand elution behaviour. This was corroborated by the fragmentationpattern, revealing an [M�H]� ion at m/z 353 and a fragment at m/z191 in the MS2 experiment. Two compounds (10, 11), whichsequentially eluted at 18.5 and 19.2 min, showed similar UV/Visspectra and an identical [M�H]� ion at m/z 337, indicating theoccurrence of p-coumaroylquinic acid isomers. According to pub-lished data (Clifford, Johnston, Knight, & Kuhnert, 2003), peaks 10and 11 were assigned to the 4- and 5-isomers of p-coumaroylqui-nic acid, producing MS2 fragments at m/z 173 and 191, respec-tively. The occurrence of chlorogenic acid and p-coumaroylquinicacid in apple seeds is consistent with previous reports (Lu & Foo,1998; Schieber et al., 2003). In the present study, however, 4-and 5-p-coumaroyl-quinic acid isomers were additionally detectedin apple seeds.

Compounds 12–16 showed UV/Vis spectral data characteristicof flavonols. Each of these compounds provided an [M�H]� ionat m/z 301 in MS2 experiments, indicating the occurrence of quer-cetin. In subsequent MS2 experiments, they showed different frag-mentation patterns due to differing saccharide moieties. Forcompounds 12 and 13, a loss of a hexose moiety (162 Da) was ob-served. By comparison of their retention times with those ofauthentic standards, compounds 12 and 13 were assigned to quer-cetin 3-O-galactoside and quercetin 3-O-glucoside, respectively.According to a loss of 146 Da, compound 16 was identified as quer-cetin 3-O-rhamnoside. This assignment was corroborated by thecomparison of its retention time with that of the correspondingreference compound. The loss of 132 Da for compounds 14 and15 upon fragmentation indicated these substances to be quercetinpentosides. Due to the lack of authentic standards, compounds 14and 15 were tentatively identified as quercetin 3-O-xyloside andquercetin 3-O-arabinoside, respectively, according to their elutionbehaviour on a comparable stationary phase as previously reported(Schieber, Hilt, Conrad, Beifuss, & Carle, 2002).

min0 10 20 30 40 50 60

mAU

0

100

200

300

400

500

600

min28 30 32 34 36 38 40 42

mAU

-100

10203040

min28 30 32 34 36 38 40 42

mAU

-100

10203040

min6 8 10 12 14 16 18 20

mAU

-10

0

10

20

min6 8 10 12 14 16 18 20

mAU

-10

0

10

20

(a)

1 2 3 45 6

7 8

8

min0 10 20 30 40 50 60

mAU

0

25

50

75

100

125

150

175

200

min12 14 16 18 20

mAU

-5

0

5

10

15

min12 14 16 18 20

mAU

-5

0

5

10

15(b)9

9

10 11

min0 10 20 30 40 50 60

mAU

-2

-1

0

1

2

3

4

5

6

(c)

12 1314

1516

Fig. 3. HPLC chromatograms of apple seed extracts at detection wavelengths of 280 nm (a), 320 nm (b) and 370 nm (c). For peak assignment see Table 1.

M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287 1283

3.3. Enzymatic oxidation of apple seed extracts

3.3.1. Analysis of the incubation media by HPLCSyntheses of the pigment POPj upon oxidation of various ASEs

and a phloridzin model solution were monitored by HPLC over aperiod of up to 32 h (Fig. 4a). Although the study was focused onthe synthesis of POPj, changes of relevant educts and intermediateswere also recorded during incubation (Fig. 4b–e), thus providingadditional information on the enzymatic reaction.

3.3.1.1. General findings. According to Fig. 4a, the ASEs (A–E) andthe phloridzin model solution (PMS) significantly differed, bothwith regard to product formation kinetics and maximum yieldsof POPj. Most distinct increases of POPj contents were observedduring the first 2 h (except for extract E), while only minor changeswere observed thereafter. This is in accordance with previous find-ings, demonstrating the conversion of POPi into POPj to be veryslow due to low affinity of mushroom PPO towards POPi (Guyotet al., 2007). Samples also showed considerable differences with

regard to the conversion of phloridzin, phloretin 20-xyloglucoside,chlorogenic acid and POPi (Fig. 4b–e). Upon oxidation, phloridzinwas converted, with POPi being simultaneously formed and accu-mulated to different extents (see Fig. 4e). POPj synthesis was prob-ably further hindered by increasingly formed oxidation products ofphenolic compounds, which were shown to significantly inhibitPPO activity (Le Bourvellec, Le Quéré, Sanoner, Drilleau, & Guyot,2004). This might be the reason why POPi was incompletely con-verted into POPj.

Interestingly, pigment formation started immediately after PPOaddition. This is in disagreement to previous studies, reporting theexistence of a lag period due to the necessary conversion of the en-zyme into its active form (Guyot et al., 2007; Oszmianski & Lee,1991). This lag phase, however, was significantly shortened bylow amounts of o-diphenolic structures, such as chlorogenic acid(Oszmianski & Lee, 1991). Such synergistic effects might explainthe missing lag phase in our oxidation experiments when applyingmore complex ASEs. Considering the model solution, the absenceof the lag phase might be ascribed to differing purities of the

Table 1Retention times, UV/Vis spectra and mass spectral characteristics of phenolic compounds in apple seed extracts.

Peak Assignment rt (min) kmax (nm) [M�H]� (m/z) HPLC-ESI(�)-MSn experiment m/z (% base peak)

1 Protocatechuic acid 6.8 260, 295 153 -MS2 [153]: 109 (100)2 Procyanidin B2 16.2 232, 280 577 -MS2 [577]: 425 (100), 407 (67), 408 (46), 451 (37), 289 (36)

-MS3 [577 ? 425]: 407 (100), 273 (19), 408 (17), 381 (9), 200 (7)3 (�)-Epicatechin 18.4 231, 278 289 -MS2 [289]: 245 (100), 205 (54), 161 (16), 246 (10), 110 (8)

-MS3 [289 ? 245]: 161 (100), 203 (47)4 Procyanidin trimer 21.6 231, 280 865 -MS2 [865]: 695 (100), 739 (50), 577 (46), 696 (41), 713 (39)

-MS3 [865 ? 695]: 543 (100), 436 (40), 587 (36), 525 (26), 677 (24)5 Phloretin dihexoside 29.5 230, 280 597 -MS2 [597]: 436 (100), 435 (43), 274 (21), 499 (19)

-MS3 [597 ? 436]: 273 (100), 274 (20)6 3-Hydroxyphloridzin 32.9 230, 284 451 -MS2 [451]: 289 (100), 290 (16), 167 (5), 354 (3), 349 (3)

-MS3 [451 ? 289]: 271 (100), 167 (99), 125 (87), 272 (29), 123 (18)7 Phloretin 20-O-xyloglucoside 34.7 230, 284 567 -MS2 [567]: 273 (100), 274 (19), 167 (6), 419 (6), 531 (3)

-MS3 [567 ? 273]: 167 (100), 168 (81)8 Phloridzin 38.8 231, 284 435 -MS2 [435]: 273 (100), 274 (14), 272 (3), 167 (2)

-MS3 [435 ? 273]: 167 (100), 168 (8), 125 (5), 123 (3)9 5-O-Caffeoylquinic acid (Chlorogenic acid) 14.0 243, 302sh, 326 353 -MS2 [353]: 191 (100), 192 (9), 179 (5), 135 (2)

-MS3 [353 ? 191]: 173 (100), 127 (67), 109 (12)10 4-O-p-Coumaroylquinic acid 18.5 232, 311 337 -MS2 [337]: 173 (100), 255 (8), 163 (8), 174 (5)

-MS3 [337 ? 173]: 93 (100), 128 (92), 137 (82)11 5-O-p-Coumaroylquinic acid 19.2 232, 311 337 -MS2 [337]: 191 (100), 163 (12), 173 (7), 192 (6)

-MS3 [337 ? 191]: 127 (100), 111 (52), 85 (35), 71 (14)12 Quercetin 3-O-galactoside 31.1 255, 354 463 -MS2 [463]: 301 (100), 300 (39), 302 (4), 381 (4), 380 (4)

-MS3 [463 ? 301]: 179 (100), 271 (64), 219 (55), 151 (49), 255 (43)13 Quercetin 3-O-glucoside 31.9 254, 355 463 -MS2 [463]: 301 (100), 300 (32), 302 (23), 179 (18), 381 (9)

-MS3 [463 ? 301]: 151 (100), 271 (38), 179 (35), 192 (31), 149 (19)14 Quercetin 3-O-xyloside 33.6 254, 356 433 -MS2 [433]: 301 (100), 302 (16), 300 (12), 331 (7), 179 (7)

-MS3 [433 ? 301]: 151 (100), 179 (85)15 Quercetin 3-O-arabinoside 35.6 261, 355 433 -MS2 [433]: 301 (100), 302 (17), 271 (7), 179 (3)

-MS3 [433 ? 301]: 230 (100), 254 (68), 301 (35), 273 (31), 152 (13)16 Quercetin 3-O-rhamnoside 36.4 255, 350 447 -MS2 [447]: 301 (100), 300 (28), 302 (18), 255 (15), 314 (12)

-MS3 [447 ? 301]: 255 (100), 107 (70), 271 (63), 179 (57), 301 (50)

Abbreviations: rt, retention time; sh, shoulder.

1284 M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287

reference compounds used. Fig. 4c illustrates that phloretin 20-O-xyloglucoside was also converted by mushroom PPO. Due to itsdihydrochalcone structure, phloretin 20-O-xyloglucoside is as-sumed to be similarly oxidised to an analogous pigment. However,since such a structure could not be detected, it was concluded thatPOPj derived from phloretin 20-O-xyloglucoside co-eluted with thePOPj structure resulting from oxidation of preponderant phlorid-zin. At the end of the experiments POPj pigment yields ranged from0.09 (extract E) to 0.71 mM (extract D), thus indicating ethanolicextracts to be most suitable for enzymatic synthesis of POPj.

3.3.1.2. Extract E. In contrast to the remaining samples, the aqueousextract E showed a completely different behaviour upon oxidation.POPj was detected only after a lag phase of about 3.5 h, but consid-erable formation was noted after �18 h, resulting in the lowestsynthesis rate (0.011 mM/h) and unsatisfactory yield (0.09 mM,27.5 h) (see Fig. 4a). Obviously, this extract contained substanceshindering pigment synthesis. These findings were underpinnedby the fact that after 4 h phloridzin (Fig. 4b) was hardly convertedinto POPi (Fig. 4e). Conversion of chlorogenic acid (Fig. 4d) andphloretin 20-O-xyloglucoside (Fig. 4c) was also negligible, indicat-ing low substrate affinity of the PPO.

3.3.1.3. Extracts A and B. Compared to the aqueous extract E, pig-ment formation was markedly enhanced for extracts A and B ob-tained by a liquid–liquid extraction of the latter. For bothextracts, synthesis rates of 0.098 mM/h and 0.071 mM/h werecomparable during the first 1.5 h, respectively. Within this phase,POPj contents steadily increased to 0.14 mM (2 h) and 0.12 mM(2 h) for extracts A and B, respectively (see Fig. 4a). Nevertheless,both extracts markedly differed with regard to the oxidation ofphloridzin into POPi. Whereas phloridzin and phloretin 20-O-xylog-lucoside were completely converted within 2 h in the case of ex-

tract B, appreciable amounts of phloridzin (0.11 mM, 26.5 h) andphloretin 20-O-xyloglucoside (0.017 mM, 26.5 h) were still deter-mined in extract A at the end of the experiment (see Fig. 4b andc). As illustrated by the peak areas in Fig. 4e, POPi rapidly accumu-lated in extract B to yield the maximum relative amount among allthe samples (1816 mAU min) within 1 h. In extract A, maximumrelative conversion was reached later being markedly lower(1192 mAU min, 2 h). The observed differences were most likelydue to the extraction conditions applied. Both extracts were ob-tained with aqueous acetone, but extract B was further purifiedby liquid–liquid extraction using ethyl acetate at pH 7. Hence, ex-tract B was almost devoid of phenolic acids, while chlorogenic acidcontents of extract A remained virtually constant during incuba-tion. Moreover, polymeric procyanidins are likely to be efficientlyremoved in extract B using ethyl acetate, as recently reported(Fromm et al., 2012a). The latter are known to significantly inhibitPPO activity with increasing average degree of polymerisation (LeBourvellec et al., 2004). Thus, in contrast to extract A which is sup-posed to contain highly polymerised procyanidins, the inhibitoryeffect of the latter on PPO activity was obviously negligible in ex-tract B (Fromm et al., 2012a). As demonstrated in a previous study(Oszmianski & Lee, 1991), the presence of chlorogenic acid must bemade responsible for the promotion of POPi conversion into POPjas observed for extract A.

3.3.1.4. Extracts C, D and phloridzin model solution. As apparent fromFig. 4a for the phloridzin model solution, extracts D and C, synthe-sis rates were highest during the first 1.5 h because the respectivePOPj formation rates of 0.492 mM/h, 0.491 mM/h, and 0.404 mM/h,respectively, were quite similar. However, considerable differenceswere observed for pigment yields, with extract D showing highestaccumulation (0.50 mM, 1.75 h). Oxidative treatment of extract C,obtained by an additional liquid–liquid extraction with ethyl

0.00

0.02

0.04

0.06

0.08

0.10

0.12

0.14

0.16

0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 5

Ch

loro

gen

ic a

cid

co

nce

ntr

atio

n (

mM

)

time (h)

0.00

0.01

0.02

0.03

0.04

0.05

0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 5

Ph

lore

tin

-2'-O

-xyl

og

lc c

on

cen

trat

ion

(m

M)

time (h)

0.0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0 0.5 1 1.5 2 2.5 3 3.5 4 4.5

Ph

lori

dzin

co

nce

ntr

atio

n (m

M)

time (h)

PO

Pj c

on

cen

trat

ion

(m

M)

time (h)

0.8

0.7

0.6

0.5

0.4

0.3

0.2

0.1

0.00 2 4 6 17 19 21 23 25 27 29 31 33

(a)

(b) (c)

(d)

D

C

E

B

APMS

DPMS

CB

A

E

EA

B

DC

E

AD

C0

200

400

600

800

1000

1200

1400

1600

1800

2000

0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 5

Are

a o

f P

OP

i (m

AU

x m

in)

time (h)

(e)

EC

D

PMS A

B

Fig. 4. Kinetics of POPj (a), phloridzin (b), phloretin 20-O-xyloglucoside (c), chlorogenic acid (d) and POPi (e) upon oxidation of apple seed extracts A–E and a phloridzin modelsolution (PMS) after adding mushroom PPO (30 U/mL). Values are means ± standard deviation (n = 2).

M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287 1285

acetate at pH 1.5, also led to the formation of high amounts of POPj(0.39 mM, 1.75 h). Pigment formation was accompanied by a rapidand complete conversion of phloridzin (Fig. 4b) and phloretin 20-O-xyloglucoside (Fig. 4c) within 15 min. Additionally, POPi was mosteffectively converted into POPj within the first 2 h, as indicated bythe marked decreases of the corresponding peak areas in Fig. 4e.

In contrast, POPj content of the phloridzin model solution wassignificantly lower (0.16 mM, 2 h), due to a considerable decreaseof the synthesis rate caused by the stagnating conversion of POPi(as shown in Fig. 4e) into POPj shortly after adding PPO (�0.5 h).Again, synergistic effects in the presence of chlorogenic acid mayhave enhanced POPj formation in comparison with the model solu-tion. Interestingly, chlorogenic acid was completely metabolisedafter 2 h, illustrating improved enzyme activity. This was probably

due to the absence of inhibiting polymeric procyanidins in extractsC and D, which were recovered using ethyl acetate and alcohol,respectively. These solvents were shown to exclude polymericprocyanidins from extraction (Fromm et al., 2012a; Guyot, Marnet,Laraba, Sanoner, & Drilleau, 1998).

3.3.2. Evaluation of colour properties upon formation of POPjL⁄, C⁄ and h� values were used to characterise colour changes of

the samples upon oxidation with PPO. Most significant changeswere observed during the first 6 h of the oxidation experiments(Fig. 5). Within this period, lightness only slightly decreased forthe phloridzin model solution as well as for extracts A, B and E,which is reflected by L⁄ values still being between 95.6 and 88.1.For extracts C and D, however, the decrease of lightness was more

78

80

82

84

86

88

90

h°v

alu

e

time (h)0 2 4 6 17 19 21 23 25 27 29 31 33

(b)

PMS

E

BA

C

D

0

20

40

60

80

100

C* v

alu

e

time (h)

0 2 4 6 17 19 21 23 25 27 29 31 33

(c)

PMSE

BA

C

D

70

75

80

85

90

95

100

L*

valu

e

time (h)0 2 4 6 17 19 21 23 25 27 29 31 33

(a)PMS

E

BA

C

D

Fig. 5. Evaluation of lightness L⁄ (a), hue angle h� (b) and chroma C⁄ (c) of apple seedextracts A–E and a phloridzin model solution (PMS) upon oxidation with mushroomPPO (30 U/mL). Values are means ± standard deviation (n = 2).

1286 M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287

pronounced, resulting in L⁄ values dropping from 90.3 and 85.2 to80.3 and 74.6, respectively. At pH 6.5, all samples exhibited h�values in a range from 83.5� to 85.5� corresponding to a yellow-orange colour. As observed for lightness, h� values generallydecreased, resulting in a shift to more reddish hues. Again, extractsC and D showed the most significant shift from 83.7� and 83.5� to81.5� and 79.9�, respectively. While lightness and hue angle de-creased, chroma values (C⁄) showed an opposite trend, correlatingwell with the formation of POPj (Fig. 4a). With proceeding pigmentformation, C⁄ values increased to different extents translating intoan increment of colour saturation and higher colour brilliance.With changes from 6.4 and 12.2 to 66.0 and 76.3, highest incre-ments were registered for extracts C and D, respectively. In con-trast, C⁄ values of the phloridzin model solution only increasedfrom 2.4 to 13.9, thus exhibiting lower colour brilliance compara-ble to that of extracts A and B. As expected, colour saturation waslowest for extract E.

Enzymatic treatment of ASEs revealed marked differences inPOPj pigment formation. Depending on the extraction procedure,synthesis rates and pigment yields significantly differed, mostlikely due to varying PPO inhibition. Compared to the phloridzinmodel solution, ASEs generally exhibited higher pigment yieldsand colour brilliance. A simple extraction with aqueous ethanol(50%, v/v) has proven to be the most effective way for the obtentionof coloured extracts from apple seeds, resulting in a maximumyield of 0.71 mM POPj (after 31.5 h). Due to its simplicity and theeasy handling of an organic solvent having relatively low toxicity,industrial scale-up may be readily performed, thus allowing thecomfortable recovery of larger quantities of such colourants. Be-sides their colouring properties such preparations may also serveas natural antioxidants, since they were demonstrated to exhibitappreciable antioxidant activities (data not shown).

The results of the present study demonstrated that apple seedsmight be fully exploited for the recovery of coloured preparationsdisplaying antioxidant activities which may represent an interest-ing alternative to synthetic dyes. Altogether, the sequential extrac-tion of highly unsaturated fatty oils (Fromm, Bayha, Carle, &Kammerer, 2012b) and antioxidant phenolic compounds fromthe defatted residual seed hulls, in particular of phloridzin as anadjuvant in the treatment of diabetes and for weight control ap-pears to be promising. In addition, the subsequent oxidation ofphloridzin derivatives, as demonstrated in this study, may providea further opportunity to perform a more economical and exhaus-tive exploitation of by-products arising from apple processing.However, before such preparations may be effectively utilised fordietary purposes, further knowledge not only on their colour sta-bility in different food matrices but most of all on their toxicityis required to rule out any health risks, since apple seeds are alsoknown to contain cyanogenic glycosides.

Acknowledgment

The authors would like to thank Mr. Christian Hüttner for hisexcellent technical assistance in performing POPj formationexperiments.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at http://dx.doi.org/10.1016/j.foodchem.2012.09.042.

References

Al-Farsi, M., & Lee, C. Y. (2008). Optimization of phenolics and dietary fibreextraction from date seeds. Food Chemistry, 108, 977–985.

Alonso-Salces, R. M., Ndjoko, K., Queiroz, E. F., Ioset, J. R., Hostettmann, K., Berrueta,L. A., et al. (2004). On-line characterisation of apple polyphenols by liquidchromatography coupled with mass spectrometry and ultraviolet absorbancedetection. Journal of Chromatography A, 1046, 89–100.

Alvarado, F., & Crane, R. K. (1962). Phlorizin as a competitive inhibitor of the activetransport of sugars by hamster small intestine, in vitro. Biochimica et BiophysicaActa, 56, 170–172.

Bellion, P., Digles, J., Will, F., Dietrich, H., Baum, M., Eisenbrand, G., et al. (2010).Polyphenolic apple extracts: Effects of raw material and production method onantioxidant effectiveness and reduction of DNA damage in caco-2 cells. Journalof Agricultural and Food Chemistry, 58, 6636–6642.

Benzie, I. F. F., & Strain, J. J. (1996). The ferric reducing ability of plasma (FRAP) as ameasure of ‘antioxidant power’: The FRAP assay. Analytical Biochemistry, 239,70–76.

Bhushan, S., Kalia, K., Sharma, M., Singh, B., & Ahuja, P. S. (2008). Processing of applepomace for bioactive molecules. Critical Reviews in Biotechnology, 28, 285–296.

Cao, X., Wang, C., Pei, H., & Sun, B. (2009). Separation and identification ofpolyphenols in apple pomace by high-speed counter-current chromatographyand high-performance liquid chromatography coupled with mass spectrometry.Journal of Chromatography A, 1216, 4268–4274.

M. Fromm et al. / Food Chemistry 136 (2013) 1277–1287 1287

Chandra, A., Rana, J., & Li, Y. (2001). Separation, identification, quantification, andmethod validation of anthocyanins in botanical supplement raw materials byHPLC and HPLC–MS. Journal of Agricultural and Food Chemistry, 49, 3515–3521.

Chinard, F. P., Taylor, W. R., Nolan, M. F., & Enns, T. (1957). Transport of glucose bythe renal tubule cells of anesthetized dogs. Science, 125, 736–737.

Clifford, M. N., Johnston, K. L., Knight, S., & Kuhnert, N. (2003). Hierarchical schemefor LC–MSn identification of chlorogenic acids. Journal of Agricultural and FoodChemistry, 51, 2900–2911.

Durkee, A. B., & Poapst, P. A. (1965). Phenolic constituents in core tissues and ripeseed of McIntosh apples. Journal of Agricultural and Food Chemistry, 13, 137–139.

Durling, N. E., Catchpole, O. J., Grey, J. B., Webby, R. F., Mitchell, K. A., Foo, L. Y., et al.(2007). Extraction of phenolics and essential oil from dried sage (Salviaofficinalis) using ethanol–water mixtures. Food Chemistry, 101, 1417–1424.

Ehrenkranz, J. R. L., Lewis, N. G., Kahn, C. R., & Roth, J. (2005). Phlorizin: A review.Diabetes/Metabolism Research and Reviews, 21, 31–38.

FAOSTAT (2010). FAO Statistical Database. Available from: http://www.fao.org[accessed 16.06.12].

Fromm, M., Bayha, S., Carle, R., & Kammerer, R. D. (2012a). Characterization andquantitation of low- and high-molecular-weight phenolic compounds in appleseeds. Journal of Agricultural and Food Chemistry, 60, 1232–1242.

Fromm, M., Bayha, S., Carle, R., & Kammerer, R. D. (2012b). Comparison of fatty acidprofiles and contents of seed oils recovered from dessert and cider apples andfurther Rosaceous plants. European Food and Research Technology, 234,1033–1041.

González-Laredo, R. F., Reyes-Navarrete, M. G., Preza y Lerma, A. M., Rosales-Castro,M., Morales-Castro, J., Gallegos-Infante, J. A., et al. (2007). Antioxidantevaluation and chemoprotection of phenolic extracts from apple seeds. Grasasy Aceites, 58, 5–9.

Goodenough, P. W., Kessell, S., Lea, A. G. H., & Loeffert, T. (1983). Mono anddiphenolase activity from fruit of Malus pumila. Phytochemistry, 22, 359–363.

Guyot, S., Marnet, N., Laraba, D., Sanoner, P., & Drilleau, J.-F. (1998). Reversed-phaseHPLC following thiolysis for quantitative estimation and characterization of thefour main classes of phenolic compounds in different tissue zones of a Frenchcider apple variety (Malus domestica var. Kermerrien). Journal of Agricultural andFood Chemistry, 46, 1698–1705.

Guyot, S., Serrand, S., Le Quéré, J. M., Sanoner, P., & Renard, C. M. G. C. (2007).Enzymatic synthesis and physicochemical characterisation of phloridzinoxidation products (POP), a new water-soluble yellow dye deriving fromapple. Innovative Food Science and Emerging Technologies, 8, 443–450.

Hwang, J. K., Kim, C. J., & Kim, C. T. (1998). Extrusion of apple pomace facilitatespectin extraction. Journal of Food Science, 63, 841–844.

Keller, P., Streker, P., Arnold, G., Schieber, A., & Carle, R. (2001). Determination ofphenolic compounds in dessert and cider apples by HPLC. Flüssiges Obst, 68,480–483.

Le Bourvellec, C., Le Quéré, J.-M., Sanoner, P., Drilleau, J.-F., & Guyot, S. (2004).Inhibition of apple polyphenol oxidase activity by procyanidins and polyphenoloxidation products. Journal of Agricultural and Food Chemistry, 52, 122–130.

Le Guernévé, C., Sanoner, P., Drilleau, J.-F., & Guyot, S. (2004). New compoundsobtained by enzymatic oxidation of phloridzin. Tetrahedron Letters, 45,6673–6677.

Lea, A. G. H. (1984). Farb- und Gerbstoffe in englischen Mostäpfeln. Flüssiges Obst, 8,356–361.

Lu, Y., & Foo, L. Y. (1997). Identification and quantification of major polyphenols inapple pomace. Food Chemistry, 59, 187–194.

Lu, Y., & Foo, L. Y. (1998). Constitution of some chemical components of apple seed.Food Chemistry, 61, 29–33.

Manzano, S., & Williamson, G. (2010). Polyphenols and phenolic acids fromstrawberry and apple decrease glucose uptake and transport by humanintestinal Caco-2 cells. Molecular Nutrition & Food Research, 54, 1773–1780.

McCann, D., Barrett, A., Cooper, A., Crumpler, D., Dalen, L., Grimshaw, K., et al.(2007). Food additives and hyperactive behaviour in 3-year-old and 8/9

year-old children in the community: A randomised, double-blinded, placebo-controlled trial. Lancet, 370, 1560–1567.

Oszmianski, J., & Lee, C. Y. (1991). Enzymatic oxidation of phloretin glucoside inmodel system. Journal of Agricultural and Food Chemistry, 39, 1050–1052.

Rezk, B. M., Haenen, G. R. M. M., van der Vijgh, W. J. F., & Bast, A. (2002). Theantioxidant activity of phloretin: The disclosure of a new antioxidantpharmacophore in flavonoids. Biochemical and Biophysical ResearchCommunications, 295, 9–13.

Rupasinghe, H. P. V., & Kean, C. (2008). Polyphenol concentrations in appleprocessing by-products determined using electrospray ionization massspectrometry. Canadian Journal of Plant Science, 88, 759–762.

Rupasinghe, H. P. V., & Yasmin, A. (2010). Inhibition of oxidation of aqueousemulsions of omega-3 fatty acids and fish oil by phloretin and phloridzin.Molecules, 15, 251–257.

Sarapuu, L., & Kheinaru, E. (1969). Oxidation of phloridzin by o-diphenol oxidase inapple buds. Physiology and Plant Biochemistry, 36, 8–17.

Schieber, A., Hilt, P., Conrad, J., Beifuss, U., & Carle, R. (2002). Elution order ofquercetin glycosides from apple pomace extracts on a new HPLC stationaryphase with hydrophilic endcapping. Journal of Separation Science, 25,361–364.

Schieber, A., Hilt, P., Streker, P., Endress, H.-U., Rentschler, C., & Carle, R.(2003). A new process for the combined recovery of pectin and phenoliccompounds from apple pomace. Innovative Food Science and EmergingTechnologies, 4, 99–107.

Schieber, A., Keller, P., & Carle, R. (2001). Determination of phenolic acids andflavonoids of apple and pear by high-performance liquid chromatography.Journal of Chromatography A, 910, 265–273.

Singleton, V. L., Orthofer, R., & Lamuela-Raventos, R. M. (1999). Analysis of totalphenols and other oxidation substrates and antioxidants by means of Folin–Ciocalteu reagent. Methods in Enzymology, 299, 152–178.

Tsao, R., Yang, R., Young, J. C., & Zhou, H. (2003). Polyphenolic profiles in eight applecultivars using high-performance liquid chromatograpy (HPLC). Journal ofAgricultural and Food Chemistry, 51, 6347–6353.

Van den Berg, R., Haenen, G. R. M. M., van den Berg, H., & Bast, A. (1999).Applicability of an improved Trolox equivalent antioxidant capacity (TEAC)assay for evaluation of antioxidant capacity measurements of mixtures. FoodChemistry, 66, 511–517.

Vanzani, P., Rossetto, M., De Marco, V., Rigo, A., & Scarpa, M. (2011). Efficiency andcapacity of antioxidant rich foods in trapping peroxyl radicals: A full evaluationof radical scavenging activity. Food Research International, 44, 269–275.

Veeriah, S., Kautenburger, T., Habermann, N., Sauer, J., Dietrich, H., Will, F., et al.(2006). Apple flavonoids inhibit growth of HT29 human colon cancer cells andmodulate expression of genes involved in the biotransformation of xenobiotics.Molecular Carcinogenesis, 45, 164–174.

Vendruscolo, F., Albuquerque, P. M., & Streit, F. (2008). Apple pomace. A versatilesubstrate for biotechnological applications. Critical Reviews in Biotechnology, 28,1–12.

Verardo, V., Bonoli, M., Marconi, E., & Caboni, M. F. (2008). Determination of freeflavan-3-ol content in barley (Hordeum vulgare L.) air-classified flours:Comparative study of HPLC-DAD/MS and spectrophotometric determinations.Journal of Agricultural and Food Chemistry, 56, 6944–6948.

Vieira, F. G. K., Borges, G. S. C., Copetti, C., Di Pietro, P. F., Nunes, E. C., & Fett, R.(2011). Phenolic compounds and antioxidant activity of the apple flesh and peelof eleven cultivars grown in Brazil. Scientia Horticulturae, 128, 261–266.

Wijngaard, H. H., & Brunton, N. (2010). The optimisation of solid–liquid extractionof antioxidants from apple pomace by response surface methodology. Journal ofFood Engineering, 96, 134–140.

Xiao, J.-S., Liu, L., Wu, H., Xie, B.-J., Yang, E.-N., & Sun, Z.-D. (2008). Rapid preparationof procyanidins B2 and C1 from Granny Smith apples by using low pressurecolumn chromatography and identification of their oligomeric procyanidins.Journal of Agricultural Food Chemistry, 56, 2096–2101.