Food Research International 54 (2013) 837–851

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Food Research International

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Co-extrusion encapsulation of canola oil with alginate: Effect of quercetinaddition to oil core and pectin addition to alginate shell on oil stability

Wei Wang, Geoffrey I.N. Waterhouse, Dongxiao Sun-Waterhouse ⁎School of Chemical Sciences, The University of Auckland, Private Bag 92019, Auckland, New Zealand

⁎ Corresponding author. Tel.: +64 9 3737599x89024;E-mail address: dx.sun-waterhouse@auckland.ac.nz (D

0963-9969/$ – see front matter © 2013 Elsevier Ltd. All rihttp://dx.doi.org/10.1016/j.foodres.2013.08.038

a b s t r a c t

a r t i c l e i n f o

Article history:Received 4 June 2013Accepted 24 August 2013

Keywords:AlginateAlginate–pectin mixturesCanola oilCo-extrusion encapsulationQuercetin

This study investigates the feasibility of encapsulating antioxidant-fortified canola oil via co-extrusion using0.67% alginate (0.67%A), 1% alginate (1%A) or high methoxyl (HM) pectin-enhanced alginate (A–P) as theencapsulant. Results show that encapsulation conditions especially the core–shell flow rates and shell wall for-mulation, influenced oil bead characteristics, core oil stability and retained phenolic content. Optical and scan-ning electron microscopy revealed that the 3 types of co-extruded oil beads were spherically shaped with theA–P beads having the largest bead size. All beads were physically and chemically robust, and remained intactafter being treated in acidified water at pH 3 for 2 h. Storage trials at 20 and 38 °C for 30 or 60 days revealedthe interplay between shell formulations and storage conditions on oil primary and secondary oxidation, hydro-lytic rancidity and total phenolic content of the encapsulated canola oils. Quercetin added to the oil core wasmore effective than vitamin E or BHT in suppressing oil oxidation at 38 °C. High performance liquid chromatog-raphy analyses indicated different decomposition pathways for quercetin in these beads during storage. FT-IRstudies confirmed the chemical composition and chemical stability of the 3 types of quercetin-containing oilbeads. 1%A and A–P shells are both acceptable and comparable for preserving quercetin-containing canola oil,with A–P and 1%A being slightly better for 30 and 60 day storage periods, respectively. Thus, it is feasible andbeneficial to deliver unsaturated oil and phenolic antioxidants in the form of pectin fibre-enhanced alginatemicrobeads.

© 2013 Elsevier Ltd. All rights reserved.

1. Introduction

Rising consumer preference for functional foods stimulates researchaimed at optimising the delivery of health-promoting substances likedietary fibres (DFs), polyphenols and unsaturated fatty acids (FAs) in di-etary forms to consumers (Kumar & Goyal, 2008; Ratnayake et al., 2000;Sherry et al., 2010; Warren et al., 2009). Unsaturated plant oils such ascanola oil are becoming an increasingly important part of the humandiet. However, these oils are highly susceptible to oxidation which ifleft unchecked leads to rancidity and an unpleasant odour and taste.

Fortifying oils with antioxidants is a well-established approach forimproving oil stability against oxidation. Vitamin E has traditionallybeen used for this purpose, although synthetic antioxidants such as bu-tylated hydroxytoluene (BHT) are no longer in favour (Kahl & Kappus,1993). Recently, more emphasis has been placed on phenolics as oil-fortifying antioxidants due to their validated health benefits and abilityto suppress lipid oxidation via donating hydrogen atoms to lipid peroxylradicals (Cheung, Szeto, & Benzie, 2007). Most phenolic antioxidants

fax: +64 9 3737422.. Sun-Waterhouse).

ghts reserved.

including quercetin can impart a bitter taste to foods (Jaeger, Axten,Wohlers, & Sun-Waterhouse, 2009). In general, consumers are unwill-ing to compromise on the taste and flavour of foods, so delivery ofhigh phenolic content foods and beverages to consumers is technicallychallenging.

Co-extrusion is a microencapsulation method that is highly suitablefor preservation of oil through creating a shell wall barrier to enclosean oil core, which itself can be fortified with an antioxidant (Fang& Bhandaria, 2010; Sun-Waterhouse, Penin-Peyta, Wadhwa, &Waterhouse, 2012; Sun-Waterhouse, Wadhwa, & Waterhouse, 2013;Sun-Waterhouse, Zhou, Miskelly, Wibisono, & Wadhwa, 2011). Suchan encapsulation approach also ameliorates undesirable taste sensa-tions such as bitterness of core substances through minimising contactwith oral taste receptors. The resultant antioxidant-fortified oil micro-beads can be used either as nutritional supplements for direct consump-tion or as a food ingredient. The shell wall matrix influences the size,shape and integrity of microbeads, as well as encapsulation efficiencyand stability of core substances (Fang & Bhandaria, 2010; Sandoval-Castilla, Lobato-Calleros, García-Galindo, Alvarez-Ramírez, & Vernon-Carter, 2010). Polysaccharides are widely used as encapsulants dueto their moderately low oxygen permeability (Sandoval-Castilla et al.,2010; Williams & Phillips, 2000). Alginate gels formed in the presence

838 W. Wang et al. / Food Research International 54 (2013) 837–851

of Ca2+ are frequently utilised as a physical barrier for microencapsula-tion because of their chemical stability, low toxicity and low immunoge-nicity (Liu et al., 2002). Pectins are health-promoting soluble fibres andoften used to increase viscosity and gel strength of food products(Thakur, Singh, & Handa, 1997). Blending different biopolymers hasproven to be a useful strategy to form shell walls with maximal protec-tion and tailored controlled release properties for core substances (Islan,de Verti, Marchetti, & Castro, 2012; Piculell, Bergfeldt, & Nilsson, 1995).Adding pectins to alginate-based encapsulant formulations may confermultiple benefits: 1) enhanced protection on core substances and 2) in-creased DF consumption and product nutritional value (Fernandez,2001). Pectins that are industrially extracted from apple pomace andcitrus peels are mostly in the form of high methoxyl (HM) pectins(N50% degree of esterification, DE) (Díaz-Rojas et al., 2004). Thus, it isof interest to examine the combined use of alginate and HM pectinsfor encapsulation of bioactive oils.

In this study, canola oil was selected as amodel unsaturated plant oilbecause of its high unsaturated FA content (i.e. monounsaturated FA62.4%, polyunsaturated FA 31.3% with 61.6% C18:1, 21.7% C18:2n−6and 9.6% C18:3n−3) (Przybylski, Mag, Eskin, & McDonald, 2005). Theeffects of co-extrusion encapsulation conditions, i.e. core–shell feedflow rates and bead hardening temperature and time were first exam-ined using the oil beads encapsulated by 1% alginate only (1%A). The op-timal encapsulation conditions were then selected to produce canola oilbeads with or without an added antioxidant encapsulated by two algi-nate formulations (0.67% sodium alginate (0.67%A) or 1%A), as well asby alginate–HM pectin solution (A–P). Quercetin was selected as amodel phenolic antioxidant for oil fortification because of its potentscavenging activity and recognised health benefits (Warren et al.,2009). The effectiveness of quercetin in preserving the encapsulatedoil was compared to that of vitamin E or BHT over 60 days storages at20 and 38 °C. Optical and scanning electron microscopy techniqueswere used to explore bead characteristics before and after treatmentat pH 3. FT-IR spectroscopy and high performance liquid chromatogra-phy (HPLC) are used to examine the compositional changes of theencapsulated oil beads during the storage trials. From this work, wehope to ascertain the best shell formulation and fortifying antioxidantfor the preservation of canola oil quality.

2. Materials and methods

2.1. Chemicals and materials

Canola oil (Simply, Malaysia) was purchased from Countdown inAuckland, New Zealand. Sodium alginate (GRINSTED® Alginate FD155,viscosity 350–550 mPa·s) was purchased from Danisco, Auckland,New Zealand. HM pectin (classic AU 201 USP, degree of esterification72–76%) was purchased from Herbstreith & Fox KG (D-75305),Neüenburg, Switzerland. Quercetin, rutin, catechin, gallic acid, BHT,α-tocopherol (vitamin E), Folin–Ciocalteu's phenol reagent (2 N),calcium chloride, active carbon, p-anisidine and tridecanoic acid werepurchased from Sigma-Aldrich Inc., St Louis, MO, USA. HPLC grade ace-tonitrile was purchased from J.T. Baker, Phillipsburg, NJ, USA. Glacialacetic acid, absolute ethanol, chloroform, concentrated HCl (36%),acetonitrile, n-hexane (95%), isooctane, methanol, potassium iodide,potassium dichromate, potassium hydrogen phthalate, sodium hydrox-ide, sodium carbonate, sodium thiosulphate and sulfuric acid werepurchased from Ajax Finechem, Auckland, New Zealand. PEG 400(carbowax™) was purchased from AMCD Australia Ltd, Mulgrave,Victoria, Australia. Ammonium chloride was purchased from May &Baker, Dagenham, England. Iodine solution was purchased fromACROS ORGANICS, New Jersey, USA. Starch soluble, sodium sulphiteand potassium chloride were purchased from AnalaR, BDH laboratorychemicals, Poole, England. Cyclohexane (analytical grade) was pur-chased from Biolab (Aust) Ltd, Scoresby, Australia. Phenolphthalein in-dicator was purchased from Scharlau Chemie (Barcelona, Spain).

2.2. Preparation of encapsulated canola oil beads

2.2.1. Preparation of encapsulant formulationsA sodium alginate solution (1%w/w)was prepared by adding the re-

quired amount of Na-alginate to pre-boiled distilledwater at 85 °C (50%of final volume), mixed at 5000 rpm for 1 min using a Silverson L5Thigh shear mixer (Silverson Machines Ltd., Chesham, Bucks, UK), andthen made up to volume with distilled water at 20 °C. The resultingsolution was then subjected to further mixing using the Silverson highshear mixer (X screen; 5000 rpm, 1 min). The alginate solution wasstored overnight at 4 °C in a fridge (SRS 535NW, Samsung, Korea) andused the following day to make beads. A 0.67%w/w alginate solutionwas prepared in the same manner.

HM pectin solution (1%w/w) was prepared by slowly adding the re-quired amount of HM apple pectin powder to distilled water of 80 °Cwith gentle stirring (speed at 3, 3001 magnetic stirring hotplate,Germany). The resultant solution was gently homogenised using theSilverson high shear mixer (X screen; 800 rpm, 1 min).

The alginate–pectin (A–P) solutionwas prepared bymixing carefullythe 1%w/walginate solutionwith the 1%w/wpectin solution at a volumeratio of 2:1, followed by gentle stirring with a magnetic stirrer (speed at3 for 2 min) to obtain a homogenous shell solution.

2.2.2. Preparation of canola oil fortified with quercetin, vitamin E or BHTQuercetin dihydrate (200 ppm, final concentration in oil) was first

mixed with PEG 400 (2%w/w, final concentration in oil) in a beaker, towhich canola oil was slowly added. The beaker containing canola oiland quercetin was then placed in an ice bath, and the mixture washomogenised using the Silverson high shear mixer (X screen;2000 rpm for 1 min followed by 5000 rpm for 2 min). Fortification ofcanola oil with BHT or vitamin E was conducted in the same mannerbut without the use of PEG. PEG was added as a plasticiser and sur-factant to help blending quercetin with oil. After preliminary trials,2%w/wwas determined to be theminimal concentration needed to dis-solve quercetin.

2.2.3. Encapsulated canola oil using co-extrusion technologyThe canola oil with or without a fortifying antioxidant was encapsu-

lated by an Inotech Encapsulator (Inotech Encapsulation AG, Dottikon,Switzerland), following the method of Sun-Waterhouse, Zhou, et al.(2011) with some modifications. During encapsulation, the core fluid(200 mL of unfortified or fortified oil in a glass syringe), and the shellfluid (i.e. 200 mL of 0.67%A, 1%A or A–P solution in a plastic syringe)were simultaneously pumped into a mixing chamber to give a core–shell fluid stream which are sprayed out through a nozzle (200 μm)under a vibration frequency of 1750 Hz and a voltage of 1.5 kV. On con-tact with a 3%/w/w CaCl2 solution (200 mL), regular-sized microbeadsformed through cross-linking. After hardening in this CaCl2 solution,the beads were collected on a 50-μm nylon mesh, washed with waterat 30 °C, and transferred to 50 mL glass tubes and filled up to a volumeof 35 mL (wrapped with aluminium foil), and freeze dried using afreeze dryer (Labconco, Total Lab Systems Ltd, New Zealand). Prelimi-nary encapsulation trials were conducted using different combinationsof core–shell feed flow rates and encapsulated bead hardeningconditions Table 1: a core feed flow rate of 30 or 60 mL/h, a shell feedflow rate of 200 and 250 mL/h, and hardening at room temperaturefor 10 min or at 4 °C for 2 h in 3% CaCl2 solution. The optimal encapsu-lation conditions were determined for preparing oil beads for the stor-age trials and pH treatment studies. The unencapsulated oil (control)was prepared by placing the canola oil in the same glass syringe (asthe core fluid), then subjecting the oil to the same encapsulation pro-cess treatment but in the absence of a shell fluid in the plastic syringe,and harvested by collection in the same beaker. The control oil wasthus exposed to the same light and heat exposures as the encapsulatedoils.

Table 1Effect of encapsulation conditions on beads encapsulated using 1% alginate.

Encapsulation conditions Bead parameters and oil loading efficiency

Shell flow rate(mL/h)

Core flow rate(mL/h)

Hardeningtime

Beads size(μm)

Wall thickness(μm)

Oil loading efficiency(%)

Breakage (%)

Before freeze drying After freeze drying

200 30 RT, 10 min 350–400 60–70 65 ± 1%ab 2 ± 0.5%b 5 ± 0.5%bc

60 400–450 40–60 62 ± 1%b 4 ± 0.5%a 8 ± 0.5%a

30 4 °C, 2 h 350–400 60–70 68 ± 1%a 2 ± 0.5%b 4 ± 0.5%c

60 400–450 40–60 65 ± 1%ab 2 ± 0.5%b 6 ± 0.5%b

250 30 RT, 10 min 450–600 70–90 62 ± 1%b 2 ± 0.5%b 5 ± 0.5%bc

60 450–600 60–80 59 ± 1%b 4 ± 0.5%a 7 ± 0.5%ab

30 4 °C, 2 h 450–600 70–90 63 ± 1%ab 4 ± 0.5%a 6 ± 0.5%b

60 450–600 60–80 60 ± 1%b 3 ± 0.5%a 6 ± 0.5%b

Note: RT = room temperature. Data are expressed as mean ± standard. a–cValues of the same column with different superscripts are significantly different (p b 0.05).

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2.2.4. Measurement of oil loading efficiency and bead breakageThe oil loading efficiency was evaluated using this equation:

Oil loading efficiency % ¼ T−Sð Þ=T� 100%

where T is the total amount of the oil (g) in syringe used for preparingbeads, and S is the amount of surface oil (g) in the CaCl2 solution.

The bead breakage was measured before and after freeze drying. Asmall amount of fresh encapsulated beads (initially suspended in a 3%CaCl2 solution) was randomly withdrawn and transferred onto a50 μm nylon mesh. An aliquot (1 g) of the beads was accuratelyweighed out, from which the broken beads were removed aided by op-tical microscopy. The remaining unbroken beads were accuratelyweighed (Wunbroken). The bead breakage before freeze drying was de-fined using the following equation: Breakage before freeze drying % =(1 − Wunbroken) × 100%.

A predetermined amount of each type of bead (30 beads) wassubjected to freeze drying. The number of the broken beads after freezedrying was assessed by optical microscopy (Section 2.6). The weightsof 30 dried beads (Wdried) and the unbroken dried beads (Wdried,unbroken)were measured. The bead breakage after freeze drying was defined as:Breakage after freeze drying % = [(W

dried− Wdried,unbroken) / Wdried] ×

100%. All determinations were run in triplicate and the mean values re-ported here.

2.3. Exposure quercetin-containing canola oil beads to acidic pH

An acidic aqueous solution was prepared by adding a 0.5% HCl solu-tion (prepared from concentrated HCl of 36.8%w/w concentration) toMilli-Q water until pH 3 (0.001 mol/L). The pH was monitored using apH metre (Schott Instrument pH Meter Lab 850, Xylem Inc, Germany).Fresh quercetin-containing beads (initially suspended in the CaCl2 solu-tion) were transferred onto a 50-μm nylon mesh, and partially dried byabsorbing excess water with a paper towel held beneath the nylonmesh. The partially dried beads (300 mg, accurately weighed)were im-mediately placed into the aqueous solution (5 mL) at pH 3 for 2 h (sim-ulating the pH of acidic food matrix and the human stomach). Gentleand regular shaking was applied during this pH treatment. After thepH treatment, thewet beadswere collected and subjected to opticalmi-croscopy examinations, with the residual aqueous solution subjected tototal phenolic content analysis by the Folin–Ciocalteu assay (to examineif any quercetin and its derived products were released from the beadsduring the pH treatment).

2.4. Storage trials

Unencapsulated canola oil and freeze dried encapsulated canola oilbeads were stored in sealed glass scintillation vials (20 mL) with thesame headspace ratio (3 mL). The bead-containing vials were flushedwith N2, tightly capped, wrapped with aluminium foil, and stored

(away from light) for 30 or 60 days at room temperature (in cooledroom, 20 ± 2 °C) or at 38 ± 1 °C (in an incubator oven, MIR 162,Sanyo Electric Co., Ltd, Osaka, Japan). Two subsamples from each vialwere withdrawn on Day 0, 30 or 60. A portion of each bead subsamplewas used for microscopic, physical and chemical analyses. Theremaining freeze dried beads in the vials were flushed with N2 andkept in a freezer (−20 °C) for later use.

2.5. Water activity of the freeze dried beads

Thewater activity (aw) of the freeze dried canola oil beads wasmea-sured at 25 °C using a dew point water activity metre (AquaLab 4TE,Decagon Devices Inc., Pullman, WA, USA). For each type of bead, dupli-cate sampleswere collected from two separate bead production batchesand triplicate measurements were performed on each of the duplicatesamples.

2.6. Microscopic examination of encapsulated oil beads

The appearance, size and wall thickness of fresh encapsulatedcanola oil beads (in one drop of 3% CaCl2 solution) were examined byoptical microscopy with a Nikon Eclipse E600 microscope (NikonCorporation, Chiyoda-ku, Tokyo, Japan) equipped with a Nikon Coolpix995 3.34 mega pixel camera (×10, Nikon corporation, Chiyoda-ku,Tokyo, Japan).

A small amount of fresh encapsulated beads (initially suspended in a3% CaCl2 solution) was transferred using a plastic dropper onto a 50 μmnylon mesh, partially dried (through using a paper towel beneath thenylon mesh to absorb excess water), and then placed directly on thesample stage of an environmental scanning electron microscope(ESEM) (FEI-Quanta-200, FEI, Hillsboro, OR, USA). The images weretaken at an electron gun accelerating voltage of 10 kV at a sample tem-perature of 2 °C and a relative humidity of 95%. Ten beads per bead pro-duction batch (for two production batches) were examined, and threepictures per bead production batch were taken.

2.7. Fourier transform infrared (FT-IR) spectroscopy

The bead ingredients (i.e. canola oil, quercetin, alginate, HM pectinandwater), aswell as the freezedried quercetin-containing oil encapsu-lated beads were examined by FT-IR spectroscopy. FT-IR absorbancespectra were collected over the range 4000–650 cm−1 (4 cm−1 resolu-tion, 32 accumulated scans) using a Perkin Elmer® Spectrum 100 FT-IRspectrometer equipped with a universal ATR sampling attachment andstandard optical geometry. A sample background was collected in airbefore sample analyses. Each ingredient or bead sample was placeddirectly onto the ATR crystal at room temperature under dim light(in room with no sunlight and direct light but reduced bulb lightingfrom 5 m away). Absorbance spectra were normalised using SigmaPlot(version 11.0).

840 W. Wang et al. / Food Research International 54 (2013) 837–851

2.8. Oil stability evaluation

2.8.1. Extraction of canola oil from the encapsulated beads and extraction ofphenolics from the obtained oil

The extraction of canola oil from encapsulated beads followed themethod of Sun-Waterhouse et al. (2012) with some modifications.Freeze dried beads (2 g) were first ground using a mortar and pestle,towhich20 mLof KCl solution (0.88%w/v)was added. After the suspen-sion was mixed using a MT19 vortex mixer (Chiltern International,Slough, UK, operating at the speed of 4), chloroform (20 mL) andmeth-anol (20 mL) were added. The resultant mixture was homogenisedusing a homogeniser (12,000 rpm for 2 min; CH 6005 model, Luzern,Switzerland). The chloroform layer was then collected, and the extrac-tion step was repeated. The two collected chloroform layers were com-bined, evaporated using a LabconcoRapidVap®Concentrator (40 min at40 °C &10 kPa; Model 79100-01, Labconco Corp., Kansas City, Missouri,USA) under N2, dried in the Ultra-Low Cold Trap Centrivap® CentrifugalConcentrator (Model 78100-01, Labconco Corp., Kansas City, MO) at40 °C for 4 h under vacuum (Sun-Waterhouse, Thakorlal, & Zhou,2011; Sun-Waterhouse, Zhou, et al., 2011; Sun-Waterhouse et al.,2012), and stored in a freezer (−80 °C) until analysis.

The extraction of the phenolics in the extracted oils followed themethod of Sun-Waterhouse, Zhou, et al. (2011)with somemodifications.An aliquot of extracted oil (0.5 g) was dissolved in n-hexane (1.25 mL),to which aqueous methanol (80% v/v, 0.5 mL) was added. The mixturewas vigorously mixed for 2 min using the MT19 vortex mixer at speed4, centrifuged at 3000 rpm for 3 min (Centrifuge 5702, Eppendorf,Hamburg, Germany), and themethanolic phase was collected and trans-ferred to a vial wrapped with aluminium foil. This methanolic extractionwas repeated three times. The combinedmethanolic extracts were driedin the Ultra-Low Cold Trap Centrivap® Concentrator at 40 °C for 4 hunder vacuum, and stored in a freezer (−80 °C) until analysis.

2.8.2. Peroxide value (PV) determinationThe PVs of the oils were determined (AOCS, 1998a) following the

procedures of Sun-Waterhouse, Thakorlal, et al. (2011), and expressedas “milliequivalent (meq) per kg of oil”.

2.8.3. p-Anisidine value (p-AV) determinationThe p-AVs of the oils were determined (AOCS, 1998b), following

the procedures of Sun-Waterhouse, Thakorlal, et al. (2011). A spectro-photometer (SpectraMax Plus 384, MDS Analytical Technologies,Hawthorn, Australia) was used to measure the absorbance at 350 nm.

2.8.4. Totox value calculationTotox values were calculated from the PVs and p-AVs of the oil sam-

ples using the equation of Totox value = 2PV + p-AV (O'Connor, Lal, &Eyres, 2007).

2.8.5. Free fatty acid (FFA) determinationFFAs were determined using the direct titration method of AOCS

(2000) and following the procedures of Sun-Waterhouse, Thakorlal,et al. (2011). FFA content was expressed as “g FFA as oleic acid per100 g oil”.

2.8.6. Iodine value (IV) determinationIV was determined using the AOCS Official Method (1997) and

expressed as “g I2/100 g oil”.

2.8.7. Analysis of total extractable phenolic contentThe total extracted phenolic content (TEPC) was determined by the

Folin–Ciocalteu assay (Singleton, Orthofer, & Lamuela-Ravento, 1997),and expressed as “mg gallic acid equivalents (GalE) per kg oil”.

2.9. High performance liquid chromatography (HPLC) phenolic profiling

An analytical HPLC (HP Agilent 1100 series, Agilent Technologies,USA) equipped with a G1313A autosampler, a G1322A degasser, aG1311AQuat pump, a G1316A column compartment and a PhenomenexLuna C18 column (4.6 × 250 nm, 5 μm particle size), was used to ana-lyse the phenolics including quercetin in the encapsulated canola oilbeads (Phani, Vinaykumar, Umamaheswara rao, & Sindhuja, 2010). Themobile phases consist of (A) 1% acetic acid (45.0%), (B) acetonitrile(15.0%) and (C)methanol (40.0%). The flow ratewas 1.0 mL/min and in-jection volume was 20 μL with a total run time of 30 min. The wave-length detection range was 200 to 540 nm.

The phenolics extracts (~1 g) obtained in Section 2.8.1 were consti-tuted in aqueous methanol (80%v/v, 0.5 mL), vortexed and centrifuged(3000 rpm for 3 min). The resultant supernatant was filtered through a0.45 μm filter (Nylon Membranes; Supelco, Bellefonte, PA, USA) andthen transferred to brown vials for HPLC analysis. The undissolved quer-cetin was recovered from the obtained residues by centrifugation andfiltration. Individual phenolics were identified based on their retentiontime and absorbance maximum (λmax). The concentrations of non-flavonoid phenolics present at 280 nm and flavonoids present at370 nmwere estimated using a catechin or quercetin external standard,respectively. Quercetin was quantified at 370 nm using internal stan-dard rutin hydrate (25 ppm). The concentration of quercetinwas calcu-lated using this equation: Concentrationquercetin = (Peak Areaquercetin ×Concentrationrutin) / Peak Arearutin.

2.10. Statistical analysis

All chemical analysis data are expressed asmean ± standard devia-tion of three replicates. Analyses were carried out using MINITAB 15(Minitab Inc., Pennsylvania, USA) statistical software with one-wayANOVA followed by Tukey's multiple comparison test at p b 0.05.

3. Results and discussion

3.1. Effect of encapsulation conditions on bead characteristics, oil loadingefficiency and breakage percentage of the 1% alginate canola oil beads

The effects of co-extrusion encapsulation conditions, in particularcore–shell feed flow rates and bead hardening temperature and time,were first examined using oil beads encapsulated with 1% alginateonly (1%A). Optical microscopy examinations (Fig. 1A–D) revealedthat all the 1%A beads prepared using different core–shell feed flowrates and hardening conditions were spherically shaped which is desir-able as itminimises the surface area to volume ratio of the beads (http://en.wikipedia.org/wiki/Surface_tension) and hence reduces O2 perme-ation through the shell to the core oil. Table 1 summaries the beadsize, wall thickness, oil loading efficiency and breakage percentage ofthe alginate encapsulated beads that were prepared using a core feedflow rate of 30 or 60 mL/h, and a shell feed flow rate of 200 and250 mL/h, and hardening at room temperature for 10 min or at 4 °Cfor 2 h in 3% CaCl2 solution. These conditionswere selected based on re-sults of our preliminary trials, and yielded individual intact beads.

The feedflow rates of core and shell solutions and the hardening timeof encapsulated beads all influenced the characteristics of oil beads. Theeffect of hardening conditions on bead hardening was consistent withthe rapid gelation and cross-linking of calcium-alginate to form the‘egg-box’ junction during co-extrusion encapsulation (Li, Yun, Xing, Liu,& Xu, 2011). At the same shell flow rate, higher core flow rates led to alarger bead size but a thinner wall thickness. This is understandable aswithin a given time period, a higher core flow rate gives a greateramount of core fluid relative to a fixed amount of shell fluid. The alginatebeads obtained at the high shell flow rate (250 mL/h) yield inconsistentbead sizes (i.e. awider bead size distribution). At the same coreflow rate,the bead size andwall thickness increased at higher shell flow rates, and

100 µm

100 µm

100 µm

100 µm

B, 30-250 mL/h A, 30-200 mL/h

C, 60-200 mL/h D, 60-250 mL/h

Fig. 1.Optical micrographs (magnification 10×) of fresh canola oil beads encapsulatedwith 1% alginate, whichwere prepared using different core–shell feed flow rates (A, 30–200 mL/h;B, 30–250 mL/h; C, 60–200 mL/h; D, 60–250 mL/h), but the same hardening conditions: at 4 °C for 2 h in 3% CaCl2 solution.

841W. Wang et al. / Food Research International 54 (2013) 837–851

the increased shell flow rate led to the presence of small individual“spherical features” in the shell (which are likely to be oil droplets orpossibly air pockets) (Fig. 1B and D). The alginate beads obtained atthe core flow rate of 60 mL/h largely tended to be present as “twins”or “triplets” and suspended as clusters in the CaCl2 solution,with unevenshell wall thickness (Fig. 1C and D). In general, these findings agreedwith the results of Whelehan and Marison (2011) who reported thatbead wall thickness was controlled by the flow rate ratio of shell tocore solutions. Room temperature was selected as it has advantages re-lating to energy cost-effectiveness. 4 °C was selected as an alternativeto RT. Our preliminary trials showed that the minimal hardening timeat RT or 4 °C was 10 min and 2 h, respectively. When the same coreflow rate and a 200 mL/h shell flow rate were used, bead hardening at4 °C for 2 h appeared to be more beneficial (i.e. higher oil loading effi-ciency) than at room temperature for 10 min. Hardening time wasfound to have no effect on bead size, a result which was also observedby Lotfipour, Mirzaeei, and Maghsoodi (2012). Longer hardening timesare expected to lead to a higher degree of cross-linking (Rosas-Ledesma, Leon-Rubio, Alarcon, Morinigo, & Balebona, 2012; Smrdel,Bogataj, & Mrhar, 2008). Deladino, Anbinder, Navarro, and Martino(2008) showed that 15 min hardening facilitated bigger beads withcomplete cross-linking and maximummechanical strength. The highestoil loading efficiencywas 68 ± 1%, and the lowest breakage percentagesbefore and after freeze dryingwere 2 ± 0.5% and 4 ± 0.5%, respectively(Table 1). The 30–200 core–shell flow rate in combination with 2 hhardening at 4 °C appeared to be the best conditions for oil encapsula-tion by alginate, because of the resultant high oil loading efficiency andlow breakage percentages before and after freeze drying. Therefore,this set of process parameters was selected as the optimal conditionsand used for all other encapsulation studies.

The breakage of some beads during freeze drying was likely due tothe fast sublimation of frozen water from the alginate gel matrix,resulting in the formation of various pores without having sufficienttime to shrink (Smrdel et al., 2008). The amount of surface oil detectedon the freezedried beadswas low and constant for the three types of en-capsulated beads in this study i.e. 0.3–0.4 g surface oil/10 g dried beads.The presence of large amounts of oil on the surface of encapsulatedbeads (surface oil) is undesirable, since the surface oil not only deterio-rates rapidly causing off-flavour but also affects the wettability anddispersability of encapsulated beads (Drusch & Mannino, 2009). Thedegree of surface wrinkling caused by freeze drying the beads wasdependent on the encapsulation parameters i.e. core and shell feedflow rates and bead hardening conditions. Most severe wrinkling wasobserved for the beads produced at a core flow rate of 60 mL/h, a shellflow rate of 200 mL/h and hardened at room temperature for 10 min.

3.2. Appearance, size and wall thickness of the quercetin-containingcanola oil beads

The freeze dried quercetin-containing canola oil beads that wereencapsulated by 0.67%A, 1%A or A–P under the optimal encapsulationconditions selected in Section 3.1. (i.e. 30–200 core–shell flow ratesand 2 h hardening at 4 °C) had water activities of 0.135 ± 0.001,0.352 ± 0.001, and 0.460 ± 0.001, respectively. The water activityvalues suggest that the freeze dried A–P beads might have a slightlyhigher risk of microbial spoilage compared to the 2 types of alginateonly beads (Sun-Waterhouse et al., 2013).

The optical and ESEM micrographs (Fig. 2) confirm the structuredintegrity of all 3 types of co-extruded canola oil beads. The ESEMmicro-graphs (Fig. 2, 1st row) show that the 0.67%A (magnification 400×),

200 µm 200 µm 300 µm

100 µm

100 µm

100 µm

100 µm 100 µm100 µm

Before pH treatmentBefore pH treatment Before pH treatment

A 0.67% Alginate B 1% Alginate C Alginate-pectin

After pH 3 for 2 h After pH 3 for 2 h After pH 3 for 2 h

Fig. 2. ESEM (1st row, before pH treatment; magnification 400× for alginate alone beads and 300× for alginate–pectin beads) and optical microscopy images (magnification 10×, before,2nd row, and after, 3rd row, the treatment at pH 3 for 2 h) of fresh (wet) canola oil beads prepared with shell formulations: A) 0.67% alginate, B) 1% alginate, and C) alginate + Highmethoxyl (HM) pectin.

842 W. Wang et al. / Food Research International 54 (2013) 837–851

1%A (magnification 400×) and A–P (magnification 300×) oil beads hadregularly spherical shape and smooth surfaces. Optical microscopyconfirmed that all the 3 types of oil beads were in spherical shapeafter encapsulation and before pH treatments (Fig. 2, 2nd row). Theaverage diameter of whole bead and oil core were: 492 ± 15 and286 ± 8 μm for 0.67%A beads, 348 ± 11 and 208 ± 6 μm for 1%Abeads, and 533 ± 16 and 367 ± 9 μm for A–P beads, respectively. TheA–P beads had the largest size. The average wall thickness decreasedin the order 0.67%A beads (~103 μm) N A–P beads (~83 μm) N 1%Abeads (~70 μm). A lower concentration of alginate resulted in signifi-cantly larger beads, although the relative size of bead to core remainedalmost constant.

When alginate is used alone, an increase of alginate concentrationwould allow more Ca2+ binding sites and a greater number of alginatestrands held together in the ‘egg-box’ structure within the bead shell,thus resulting in a higher degree of cross-linking and a more rigid andcompact matrix (Liu et al., 2002; Mandal, Kumar, Krishnamoorthy, &Basu, 2010). Accordingly, an elevated alginate concentration increasedthe viscosity of alginate gel and subsequently a slower diffusion of coresubstances, such as small polar phenolic compounds that were poten-tially derived from quercetin decomposition at pH like 3 (seeSection 3.3), through the alginate ‘egg-box’ cavities to the outer solution(Liu et al., 2002; Mandal et al., 2010). When the shell was A–P mixture,the actual concentration of alginate in A–P was 0.67% (as a result ofmixing 1% alginate solution with 1% pectin solution at a 2:1 volumeratio). Thus, it is not surprising that the shell wall thickness of the A–Pbeads ranked in between those of the 0.67%A and 1%Abeads. Biopolymercomposition was reported to influence bead diameter and roundness(Sandoval-Castilla et al., 2010). Based on microscopy observations, theA–P combination was suitable for encapsulation. Alginate and pectin

are both polyuronates, and the Ca2+ binding mechanism occurred inboth alginate and the unmethoxylated pectins in HM pectin (althoughN50% methoxylated pectins in HM pectin did not gel relying on Ca2+

but sugar or acid) (Siew & Williams, 2005; Whistler & BeMiller, 1997).Madziva, Kailasapathy, and Phillips (2005) reported that A–P encapsu-lated beads possessed greater encapsulation efficiency, but less sphericalshape than the alginate encapsulated beads. Islan et al. (2012) reportedthat incorporating 1% HM pectin into 2% alginate resulted in less rigidstructure of microspheres than those prepared using alginate only.Thus, it is expected that the HMpectin component influences themicro-structure and network density of encapsulated beads (Morris & Chilvers,1984; Sandoval-Castilla et al., 2010).

3.3. Effect of acidic treatment on the quercetin-containing oil beads

The integrity of the quercetin-containing 0.67%A, 1%A andA–P beadswas retained after the acidic treatment at pH 3 for 2 h (Fig. 2, 3rd row),suggesting that these bead shells were all chemically and structurallyrobust. The size of these beads, however, decreased significantly afterthe pH treatment: The 1%A beads had the smallest bead size reduction(12.9%), whilst the 0.67%A and A–P beads exhibited comparable size re-duction (37.6% and 29.1%, respectively). This shrinkage enabled a morecompact molecular arrangement within the shell network. Previousstudies have reported that alginate beads tended to shrink in highlyacidic media (Rayment et al., 2009), due to either strong hydrogen in-teractions or intermolecular lactonisation (Tataru, Popa, & Desbrieres,2011). In this study, the oil beads did not rupture at low pH.

The total phenolics (including quercetin and its derived products)released from the beads into the acidic water were evaluated bythe Folin–Ciocalteu assay (Table 2), and decreased in the order

Table 2Total phenolic content of the aqueous solutions obtained following the treatments ofquercetin-containing oil beads at pH 3 for 2 h.

Sample Total phenolic content(μg gallic acid equivalent per mLsolution or mg beads)

Water solution for pH treatment 0.175 ± 0.001c

0.67% Alginate 0.238 ± 0.001a

1% Alginate 0.213 ± 0.001b

Alginate-pectin 0.207 ± 0.001bc

Note: Data are expressed as mean ± standard. a–cvalues of the same column withdifferent superscripts are significantly different (p b 0.05).

843W. Wang et al. / Food Research International 54 (2013) 837–851

0.67%A N 1%A N A–P beads. The release characteristics of bead shells arestrongly dependent on their permeability (which was affected by thepolarity, density, porosity, homogeneity and thickness of shell wall ma-terials) (Donhowe & Fennema, 1993). Alginate is pH sensitive (Whistler& BeMiller, 1997) rendering it suitable for the intestinal delivery sys-tems. Alginate solutions are chemically stable at pH 5.5–10 at roomtemperature (McDowell, 1977). When pH N pKa, the carboxylategroups are ionised in solution and alginate polymer acts like a polyelec-trolyte. When pH b pKa, the carboxyl groups remain unionised andalginate polymer behaves more like a neutral polymer depending onthe ratio of β-D-mannuronic acid (M) and α-L-guluronic (G) acid(Bergfeldt, Piculell, & Tjerneld, 1995). When pH b 4, alginate isconverted to alginic acid forming a viscous acid gel (Rayment et al.,2009), thus retarding the loss of encapsulated phenolics. Alginate con-centration also affects the porosity of its gel matrix (Goh, Heng, &Chan, 2012). Blandino, Macías, and Cantero (2001) found that capsulesprepared with 1% sodium alginate were more robust than those withlower alginate concentrations, because a more densely cross-linkedgel structure could be formed. Thus, it is not surprising that a slightlygreater amount of phenolics were released from the 0.67%A beadsthan the 1%A beads.

Mixing alginate with HMpectin modified the characteristics of beadshells and the release of substances from bead cores (Madziva et al.,2005; Sun-Waterhouse et al., 2012). The gelation of pure HM pectin in-volves various intermolecular interactions, with the junction zonesbeing stabilised by both hydrogen bonds and hydrophobic interactionsinvolving ester methyl groups (Oakenfull & Scott, 1984; Walkinshaw& Arnott, 1981). In pure HM pectin, the proportion of the ionisedcarboxyl groups present is smaller (b50%) than the proportion of meth-ylated carboxylate groups. Thus, only a small number of hydrogenbonds are formed with stronger interactions like ion-dipole and gener-ation of electrostatic complexes between anionic alginate and pectinbeing dominant. The addition of HM pectin naturally reduces the elec-trostatic repulsion and permits molecular association with alginate car-boxyls in free acid form and facilitates subsequent gelling (Morris &Chilvers, 1984). The interaction between pectin and alginate in mixedgels is a heterogeneous association between the poly-G blocks of algi-nate and the methyl ester regions of pectin of low charge, packing to-gether in rigid ribbons (Voo, Ravindra, Tey, & Chan, 2011). The pH iscritical to the gel formation for pectins especially HM pectins. Low pHsincrease the percentage of unionised carboxyl groups, thus reducingelectrostatic repulsion between adjacent pectin chains (Whistler &BeMiller, 1997). The pH levels of the 1%A solution and 0.67%A solutionwere ~7.2 and ~6.9, respectively. Pectin is a weak acid with a pKa

value of ~4 and 1% HM pectin solution has a pH ~3.5. When 1%A solu-tion was mixed with 1% HM pectin solution at a volume ratio of 2:1for encapsulation, the pH of the mixed shell solution was ~5.6. Whenthe A–P beads were exposed to acidified water at pH 3, the HM pectincomponent in the A–P shell should be still stable (because HM pectingenerally has excellent stability at all temperatures at pH of 2.5–4.5)(Oakenfull & Scott, 1984;Whistler & BeMiller, 1997). The alginate com-ponent in the A–P shell, however, shrank and became more viscous

(Rayment et al., 2009). The different responses to pH of alginate andHM pectin are expected to influence shell porosity and density, and beresponsible for different degrees of loss of phenolics from the encapsu-lated beads.

3.4. Stability of core oil and total extracted phenolic content (TEPC) in beadsafter storages

The degree of oxidation (evaluated based on PV, p-AV and Totoxvalues), extent of hydrolytic rancidity (evaluated based on FFA con-tents), change in unsaturation (evaluated based on IV values), and reten-tion of phenolics (evaluated based on TEPC values) of the encapsulatedcanola oil beads over 60 day storage trials were summarised in Table 3(at 20 °C) and Table 4 (at 38 °C).

The PVs of each encapsulated oil increased with storage time, and theincrease was greater at 38 °C (Tables 3 and 4). Such PV increases weredue to a higher rate for hydroperoxide formation rather than decomposi-tion (Lee, Lee, & Choe, 2007). For each type of beads, addition of an anti-oxidant (i.e. quercetin, vitamin E or BHT) mostly reduced primary oiloxidationbut exceptionswere found especially in thepresence of vitaminE The effectiveness decreased in the order quercetin N BHT N vitamin Eat either 20 or 38 °C. After 60 days at 20 °C, the quercetin-containing en-capsulated oils had a similar PV (9 meq/kg oil) which was lower thanthat of control oil (unencapsulated oil, 13.4 meq/kg oil) (Table 3). After60 days at 38 °C (Table 4), the PV of control oil was the highest(16.5 meq/kg oil), while the quercetin-containing oil encapsulated by1%A had a slightly lower PV (10.1 meq/kg oil) than that encapsulatedby 0.67%A or A–P (~12.5 meq/kg oil).

The p-AVs of the control and encapsulated oils increased after60 days at 20 °C or 38 °C, indicating an increased concentration of sec-ondary oil oxidation products. At either storage temperature, little dif-ference was detected in p-AV among the 3 types of antioxidant foreither the 0.67%Abeads or the 1%A beads. But for theA–P beads, vitaminEwas slightlymore effective than BHT andquercetin in suppressing sec-ondary oil oxidation, whichmay be associated with polarity differencesbetween these antioxidants and oil primary oxidation products. After60 days and in the presence of quercetin, the 1%A beads had the lowestp-AVs (2.20 and 2.98 at 20 °C or 38 °C, respectively).

Totox values estimate the overall oil oxidation status. After 60 days at20 °C, the control oil had a higher Totox value (29.4, close to the legallyacceptable value 30, O'Connor et al., 2007) than the quercetin-containingencapsulated oils (20–22). After 60 days at 38 °C, the Totox valuedecreased in the order control oil (36.9) N quercetin-containing A–P(30.1) N quercetin-containing 0.67%A (27.4) N quercetin-containing1%A (23.2). After 30 days at either temperature and in the presence ofquercetin, the A–P beads had lower Totox values compared to theother 2 types of beads. Thus, in the presence of quercetin, A–P mayform the best shell against oil oxidation for a short storage period (i.e.30 days), but 1%A appeared to be the best shell for 60 day storageperiods.

The FFA of canola oil was low on Day 0 (0.08%) but increased afterstorage, suggesting moisture capture and moisture permeability. Inthe absence of an added antioxidant, the encapsulated oils had compa-rable FFA contents after 60 days storage. No discernable differences insuppressing oil hydrolytic rancidity were observed among the 3 typesof antioxidants i.e. quercetin, vitamin E and BHT. The FFA contentsshowed fluctuation due to the nature of the lipid and its surroundingenvironment (Hung & Slinger, 1981). After storage at 20 °C and 38 °C,the differences among the FFA readings of different encapsulated oilsunder the same storage conditions were insignificant.

Iodine value (IV) indicates unsaturation status of lipids thus provid-ing an estimate of oil nutritional value (i.e. unsaturated oil is generallyconsidered good for health) and oil stability (i.e. unsaturated lipids aremore susceptible to rancidity) (Toscano, Riva, Pedretti, & Duca, 2012).IV of oil typically decreases with storage time before levelling off, be-cause oxidation decreases oil unsaturation via abstraction of hydrogen

Table 3Peroxide value, p-anisidine value, free fatty acid content, iodine value and total extracted phenolic content of control and encapsulated oils after storage at room temperature.

Oil sample Storage (day) PV (meq/kg oil) p-AV Totox FFA (g/100 g oil) IV (g I2/100 g oil) TEPC (mg GalE/kg oil)

Control 0 5.0 ± 0.1C,a 1.6 ± 0.1C,de 11.6 ± 0.2C,b 0.1 ± 0.1B,a 185 ± 0.5A,ab 21.2 ± 0.1B,e

30 9.7 ± 0.3B,a 1.9 ± 0.1B,d 21.2 ± 0.5B,a 0.2 ± 0.1B,a 180 ± 0.5A,b 23.0 ± 0.8B,c

60 13.4 ± 0.2A,b 2.7 ± 0.1A,de 29.4 ± 0.3A,b 0.4 ± 0.1A,a 176 ± 0.4B,a 26.9 ± 0.4A,d

A′ + O 0 4.9 ± 0.1C,a 2.3 ± 0.1B,b 12.1 ± 0.1C,a 0.1 ± 0.1B,a 140 ± 0.3B,d 24.2 ± 0.2B,d

30 9.3 ± 0.6B,a 2.2 ± 0.1B,c 20.9 ± 0.8B,a 0.2 ± 0.1B,a 179 ± 0.3A,b 18.8 ± 0.6C,d

60 13.1 ± 0.3A,b 2.9 ± 0.1A,cd 29.1 ± 0.5A,b 0.4 ± 0.1A,a 138 ± 0.3B,d 29.5 ± 0.6A,cd

A′ + O + Q 0 4.8 ± 0.1C,a 2.6 ± 0.1B,a 12.2 ± 0.2C,a 0.1 ± 0.1A,a 168 ± 0.1B,c 36.1 ± 0.5A,b

30 6.9 ± 0.1B,c 2.8 ± 0.1B,a 16.6 ± 0.1B,b 0.2 ± 0.1A,a 180 ± 0.5A,b 23.5 ± 0.3B,c

60 9.0 ± 0.7A,e 4.0 ± 0.2A,a 22.0 ± 0.9A,cd 0.2 ± 0.1A,a 135 ± 0.9C,d 32.6 ± 0.1A,cd

A′ + O + VE 0 4.9 ± 0.1C,a 1.7 ± 0.1C,d 11.5 ± 0.1C,b 0.1 ± 0.1B,a 178 ± 0.8A,b 30.8 ± 0.1B,c

30 8.4 ± 0.6B,b 2.3 ± 0.1B,c 19.9 ± 0.9B,a 0.2 ± 0.1AB,a 169 ± 0.4B,bc 26.9 ± 0.3C,b

60 12.7 ± 0.5A,b 3.5 ± 0.1A,b 28.9 ± 0.7A,b 0.3 ± 0.1A,a 112 ± 0.5C,f 46.5 ± 0.8A,a

A′ + O + BHT 0 4.8 ± 0.1C,a 1.9 ± 0.1C,c 11.4 ± 0.2C,b 0.1 ± 0.1B,a 170 ± 0.9A,bc 31.4 ± 0.1B,c

30 7.6 ± 0.3B,bc 2.2 ± 0.1B,c 17.4 ± 0.5B,ab 0.2 ± 0.1AB,a 158 ± 0.7B,c 30.8 ± 0.6B,ab

60 11.0 ± 0.2A,c 3.4 ± 0.1A,b 25.2 ± 0.3A,c 0.3 ± 0.1A,a 110 ± 0.9C,f 40.3 ± 0.4A,ab

A + O 0 4.8 ± 0.1C,a 1.3 ± 0.1B,f 10.9 ± 0.2C,b 0.1 ± 0.1B,a 177 ± 0.5A,b 25.2 ± 0.1A,d

30 7.9 ± 0.1B,b 2.0 ± 0.1A,cd 17.8 ± 0.2B,ab 0.2 ± 0.1AB,a 176 ± 0.4A,bc 26.2 ± 0.6A,b

60 12.8 ± 0.1A,b 2.2 ± 0.1A,ef 27.8 ± 0.1A,bc 0.4 ± 0.1A,a 172 ± 0.3B,ab 22.5 ± 0.5B,e

A + O + Q 0 4.9 ± 0.1C,a 1.2 ± 0.1C,g 12.0 ± 0.1C,a 0.1 ± 0.1A,a 176 ± 0.3A,b 48.7 ± 0.1A,a

30 7.0 ± 0.3B,c 1.8 ± 0.1B,de 15.9 ± 0.4B,bc 0.2 ± 0.1A,a 171 ± 0.3A,bc 32.3 ± 0.3C,a

60 9.0 ± 0.2A,d 2.2 ± 0.1A,ef 20.2 ± 0.3A,cd 0.2 ± 0.1A,a 159 ± 0.5C,b 37.8 ± 0.4B,b

A + O + VE 0 4.8 ± 0.1C,a 1.5 ± 0.1C,e 11.1 ± 0.2C,b 0.1 ± 0.1A,a 187 ± 0.1A,a 28.2 ± 0.2B,cd

30 7.9 ± 0.5B,b 2.1 ± 0.1B,cd 17.9 ± 0.7B,ab 0.2 ± 0.1A,a 162 ± 0.4B,c 26.0 ± 0.2C,b

60 12.2 ± 0.3A,bc 2.5 ± 0.1A,e 26.9 ± 0.5A,bc 0.2 ± 0.1A,a 125 ± 0.7C,e 43.2 ± 1.4A,a

A + O + BHT 0 4.9 ± 0.3C,a 1.5 ± 0.1B,e 11.3 ± 0.4C,b 0.1 ± 0.1B,a 130 ± 0.1A,de 32.3 ± 0.6B,bc

30 7.6 ± 0.4B,bc 1.7 ± 0.1A,de 16.9 ± 0.5B,b 0.2 ± 0.1AB,a 127 ± 0.1AB,d 26.8 ± 0.8C,b

60 11.2 ± 0.3A,c 1.9 ± 0.1A,f 24.3 ± 0.4A,c 0.3 ± 0.1A,a 121 ± 0.9B,e 35.1 ± 1.8A,bc

A + P + O 0 4.9 ± 0.1C,a 1.7 ± 0.1B,d 11.5 ± 0.2C,b 0.1 ± 0.1B,a 131 ± 0.5C,de 23.2 ± 0.5B,de

30 8.0 ± 0.7B,b 1.9 ± 0.1B,d 17.8 ± 1.1B,ab 0.2 ± 0.1B,a 187 ± 0.1A,a 23.5 ± 0.2B,c

60 14.3 ± 0.6A,a 2.3 ± 0.1A,e 30.9 ± 0.9A,a 0.4 ± 0.1A,a 155 ± 0.4B,bc 25.9 ± 0.4A,de

A + P + O + Q 0 4.8 ± 0.1C,a 2.5 ± 0.1B,a 12.0 ± 0.2C,a 0.1 ± 0.1A,a 126 ± 0.5B,e 35.0 ± 0.2A,b

30 5.5 ± 0.3B,d 2.5 ± 0.1B,b 13.4 ± 0.4B,c 0.2 ± 0.1A,a 178 ± 0.7A,b 23.0 ± 0.2C,c

60 9.0 ± 0.3A,d 3.1 ± 0.1A,c 21.1 ± 0.4A,cd 0.2 ± 0.1A,a 171 ± 0.5A,ab 31.8 ± 0.5B,c

A + P + O + VE 0 4.9 ± 0.1C,a 1.5 ± 0.1B,e 11.3 ± 0.2C,b 0.1 ± 0.1B,a 174 ± 0.9A,b 28.9 ± 0.3A,cd

30 8.1 ± 0.4B,b 1.6 ± 0.1B,e 17.8 ± 0.6B,ab 0.1 ± 0.1B,a 159 ± 0.1B,c 26.3 ± 0.3B,b

60 13.3 ± 0.7A,b 2.8 ± 0.1A,d 29.4 ± 1.0A,b 0.3 ± 0.1A,a 148 ± 0.1C,c 29.8 ± 0.4A,cd

A + P + O + BHT 0 4.7 ± 0.3C,a 1.7 ± 0.1B,d 11.1 ± 0.4C,b 0.1 ± 0.1B,a 172 ± 0.1A,bc 28.1 ± 0.1C,cd

30 6.8 ± 0.5B,c 1.8 ± 0.1B,de 15.4 ± 0.7B,bc 0.1 ± 0.1B,a 133 ± 0.3B,cd 32.3 ± 0.5B,a

60 11.0 ± 0.7A,c 3.6 ± 0.1A,b 25.6 ± 0.9A,c 0.3 ± 0.1A,a 115 ± 0.9C,f 38.4 ± 0.3A,b

Note: PV = peroxide value, p-AV = p-anisidine value, FFA free = fatty acid, IV = iodine value, TEPC= total extracted phenolic content, GalE = gallic acid equivalents, A = 1% alginate,A′= 0.67% alginate, O = canola oil, Q = quercetin, vitamin E = VE, and P = High methoxyl pectin. A–Cvalues of the same analysis for the same oil sample but different storage periods(Days 0, 30 and 60) with different superscripts are significantly different (p b 0.05). a–fValues of the same analysis and on the same storage day but for different oil samples with differentsuperscripts are significantly different (p b 0.05).

844 W. Wang et al. / Food Research International 54 (2013) 837–851

atoms adjacent to a double bond and leads to formation of free radicalsuntil complete oxidation of the accessible double bonds in FAs (Naz,Sheikh, Siddiqi, & Sayeed, 2004). In this study, the IV readings of thecontrol and encapsulated oils generally decreased after storage. After60 days at 20 °C, the quercetin-containing A–P beads had the highestIV (171 ± 0.5 g I2/100 g oil). The IV of 1%A and 0.67%A beads were159 ± 0.5 and 135 ± 0.9 g I2/100 g oil, respectively. After 60 days at38 °C, the control oil had lower IV (126 ± 0.1 g I2/100 g oil) comparedto the 3 types of quercetin-containing encapsulated oils. The IV of 1%Abeads (150 ± 0.1 g I2/100 g oil) was higher than those of the A–P and0.67%A beads (143 ± 0.4 and 140 ± 0.3 g I2/100 g oil, respectively).Quercetin generally exerted better preservation of unsaturated FAs inthe encapsulated oils compared to vitamin E or BHT. Some of the Day30 IV readings of encapsulated oilswere relatively high,whichmight re-sult from interferents such as aldehydes and ketones newly formeddur-ing oil oxidation.

As expected, the TEPC values were higher for the beads with anadded antioxidant. Different TEPC patterns were found for the oil beadsencapsulated with different types of shell. In presence of quercetin, the1%A beads had a slightly higher TEPC than the 0.67%A and A–P beadsafter 30 or 60 days at 20 °C. However, at 38 °C, the 1%A beads exhibiteda much lower TPEC (32.6 ± 0.2 mg GalE/kg oil) compared with theother 2 types of beads (53.7 ± 0.4 and 54.5 ± 0.4 mg GalE/kg oil)after 60 days, whilst the A–P beads had a slightly higher TEPC (40.8 ±

0.7 mg GalE/kg oil) than the 0.67%A and 1%A beads (33.0 ± 0.4 and38.5 ± 0.3 mg GalE/kg oil, respectively) after 30 days. The TEPCdetected by the Folin–Ciocalteu assay after extraction represents the ex-tractable phenolic compounds present in the oil beads. Factors such asbead matrix, storage temperature and time, and extraction method allinfluence TEPC values (Sun-Waterhouse et al., 2012). The amounts ofextracted canola oil from the 0.67%A, 1%A and A–P beads were 86.6,78.3 and 52.5 g/100 dried bead, respectively. The TEPC values were thenet results of self-degradation of antioxidants (especially at elevatedstorage temperatures or prolonged storage times or both), the consump-tion of added antioxidants for protecting oil against deterioration, andthe different extractability of antioxidant from core canola oils encapsu-lated by different bead shell matrices.

Interplay exists between shell formulations and storage conditionswhich ultimately influence the detected PV, p-AV, FFA, IV and TEPCreadings. There was a close correlation between the TEPC values andthe status of oil deterioration, indicating that a significant portion ofadded antioxidants were used to suppress oil rancidity. Oil stability de-pends on chemical and physical factors such as light, temperature, pH,FA composition and available oxygen (Frankel, Satué-Gracia, Meyer, &German, 2002). BHT and tocopherols both contain phenolic compo-nents and are lipophilic organic compounds. Quercetin is quite polarbut has low water solubility (Huber, Rupasinghe, & Shahidi, 2009),and in this study, was distributed in oil with the aid of PEG.

Table 4Peroxide value, p-anisidine value, free fatty acid content, iodine value and total extracted phenolic content of control and encapsulated oils after storage at 38 °C.

Oil sample Storage (day) PV (meq/kg oil) p-AV Totox FFA (g/100 g oil) IV (g I2/100 g oil) TEPC (mg GalE/kg oil)

Control 0 5.0 ± 0.1C,a 1.6 ± 0.1C,bc 11.6 ± 0.2C,b 0.1 ± 0.1B,a 185 ± 0.7A,a 21.2 ± 0.1B,e

30 11.3 ± 0.2B,a 2.1 ± 0.1B,e 24.8 ± 0.3B,a 0.3 ± 0.1A,a 170 ± 0.5B,ab 19.6 ± 0.2C,f

60 16.5 ± 0.1A,b 3.9 ± 0.1A,bc 36.9 ± 0.1A,a 0.2 ± 0.1AB,a 126 ± 0.1C,bc 26.1 ± 0.3A,ef

A′ + O 0 4.9 ± 0.1C,a 2.3 ± 0.1C,ab 12.1 ± 0.1C,a 0.1 ± 0.1B,a 139 ± 0.5C,de 24.2 ± 0.3B,de

30 10.1 ± 0.7B,b 2.7 ± 0.1B,c 22.9 ± 0.9B,ab 0.3 ± 0.1A,a 162 ± 0.5A,bc 23.1 ± 0.3B,e

60 15.7 ± 0.3A,bc 3.3 ± 0.1A,cd 34.7 ± 0.4A,ab 0.2 ± 0.1AB,a 133 ± 0.5B,b 27.1 ± 0.1A,ef

A′ + O + Q 0 4.8 ± 0.1C,a 2.6 ± 0.1B,a 12.2 ± 0.2C,a 0.1 ± 0.1A,a 167 ± 0.1A,c 31.5 ± 0.3B,c

30 8.9 ± 0.1B,c 2.4 ± 0.1B,d 20.3 ± 0.1B,b 0.2 ± 0.2A,a 166 ± 0.5A,b 33.0 ± 0.4B,d

60 12.1 ± 0.1A,e 3.2 ± 0.1A,d 27.4 ± 0.2A,bc 0.2 ± 0.1A,a 140 ± 0.3B,ab 53.7 ± 0.4A,c

A′ + O + VE 0 4.9 ± 0.1C,a 1.7 ± 0.1C,b 11.5 ± 0.1C,b 0.1 ± 0.1A,a 169 ± 0.3A,c 26.8 ± 0.1C,cd

30 9.7 ± 0.1B,bc 2.6 ± 0.1B,cd 21.9 ± 0.2B,b 0.2 ± 0.1A,a 140 ± 0.4B,e 50.3 ± 0.4B,b

60 15.5 ± 0.1A,bc 3.5 ± 0.1A,cd 34.5 ± 0.2A,ab 0.2 ± 0.1A,a 112 ± 0.5C,cd 60.2 ± 0.6A,bc

A′ + O + BHT 0 4.8 ± 0.1C,a 1.9 ± 0.1C,b 11.4 ± 0.2C,b 0.1 ± 0.1B,a 168 ± 0.3A,c 27.0 ± 0.2B,cd

30 8.3 ± 0.1B,cd 2.3 ± 0.1B,de 18.8 ± 0.2B,c 0.3 ± 0.1A,a 123 ± 0.4B,de 66.8 ± 0.3A,a

60 13.5 ± 0.6A,cd 3.1 ± 0.2A,cd 30.1 ± 0.9A,b 0.3 ± 0.1A,a 118 ± 0.4C,cd 67.9 ± 0.4A,b

A + O 0 4.8 ± 0.1C,a 1.3 ± 0.1C,c 10.9 ± 0.2C,b 0.1 ± 0.1B,a 177 ± 0.2A,b 25.2 ± 0.1B,d

30 8.1 ± 0.2B,d 2.8 ± 0.1B,c 18.9 ± 0.2B,c 0.3 ± 0.1A,a 175 ± 0.2A,a 24.1 ± 0.3B,e

60 13.8 ± 0.3A,cd 3.3 ± 0.1A,cd 30.9 ± 0.4A,b 0.3 ± 0.1A,a 142 ± 0.2B,ab 28.3 ± 0.1A,e

A + O + Q 0 4.9 ± 0.1C,a 1.2 ± 0.1C,c 12.0 ± 0.1C,a 0.1 ± 0.1B,a 176 ± 0.3A,b 48.7 ± 0.1A,a

30 8.7 ± 0.3B,cd 2.4 ± 0.1B,d 19.7 ± 0.4B,bc 0.2 ± 0.1AB,a 151 ± 0.2B,c 38.5 ± 0.3B,cd

60 10.1 ± 0.2A,f 3.0 ± 0.1A,d 23.2 ± 0.2A,c 0.3 ± 0.1A,a 150 ± 0.1B,a 32.6 ± 0.2C,d

A + O + VE 0 4.8 ± 0.1C,a 1.5 ± 0.1C,bc 12.1 ± 0.2C,a 0.1 ± 0.1A,a 182 ± 0.1A,a 26.6 ± 0.2C,cd

30 9.6 ± 0.7B,bc 2.5 ± 0.1B,cd 21.7 ± 0.9B,b 0.2 ± 0.1A,a 126 ± 0.7B,de 50.0 ± 0.4B,b

60 13.7 ± 0.9A,cd 3.7 ± 0.1A,c 31.1 ± 1.2A,b 0.2 ± 0.1A,a 118 ± 0.7C,cd 74.9 ± 1.4A,a

A + O + BHT 0 4.9 ± 0.3C,a 1.5 ± 0.1C,bc 11.3 ± 0.4C,b 0.1 ± 0.1B,a 130 ± 0.1A,de 30.2 ± 0.6C,c

30 8.2 ± 0.6B,d 2.2 ± 0.1B,e 18.7 ± 0.8B,c 0.2 ± 0.1AB,a 118 ± 0.7B,e 49.7 ± 0.1B,b

60 12.6 ± 0.4A,de 3.4 ± 0.1A,cd 28.6 ± 0.5A,bc 0.3 ± 0.1A,a 113 ± 0.6C,cd 73.1 ± 0.4A,a

A + P + O 0 5.0 ± 0.1C,a 1.7 ± 0.1C,b 11.5 ± 0.2C,b 0.1 ± 0.1B,a 132 ± 0.4C,de 23.2 ± 0.5B,de

30 7.9 ± 0.4B,de 3.4 ± 0.1B,a 19.1 ± 0.6B,c 0.3 ± 0.1A,a 178 ± 0.7A,a 22.4 ± 0.5B,ef

60 16.0 ± 0.5A,b 4.5 ± 0.1A,ab 36.5 ± 0.7A,a 0.2 ± 0.1AB,a 145 ± 0.2B,ab 24.6 ± 0.2A,f

A + P + O + Q 0 4.8 ± 0.1C,a 2.5 ± 0.1C,a 12.0 ± 0.2C,a 0.1 ± 0.1B,a 126 ± 0.4C,e 38.8 ± 0.2C,b

30 7.2 ± 0.5B,e 3.0 ± 0.1B,b 17.4 ± 0.6B,d 0.2 ± 0.1AB,a 149 ± 0.4A,c 40.8 ± 0.7B,c

60 13.1 ± 0.7A,d 3.9 ± 0.1A,bc 30.1 ± 0.9A,b 0.3 ± 0.1A,a 143 ± 0.4B,ab 54.5 ± 0.4A,c

A + P + O + VE 0 5.0 ± 0.1C,a 1.5 ± 0.1C,bc 11.3 ± 0.2C,b 0.1 ± 0.1B,a 155 ± 0.9A,d 32.1 ± 0.3B,bc

30 9.1 ± 0.6B,c 2.2 ± 0.1B,e 20.3 ± 0.9B,b 0.2 ± 0.1AB,a 128 ± 0.6B,de 33.4 ± 0.4B,d

60 17.3 ± 0.1A,a 4.3 ± 0.1A,b 38.9 ± 0.1A,a 0.3 ± 0.1A,a 121 ± 0.7C,c 55.0 ± 0.9A,c

A + P + O + BHT 0 4.7 ± 0.3C,a 1.7 ± 0.1C,b 11.1 ± 0.4C,b 0.1 ± 0.1B,a 165 ± 0.1A,c 30.6 ± 0.6B,c

30 8.9 ± 0.4B,c 2.7 ± 0.1B,c 20.4 ± 0.5B,b 0.2 ± 0.1AB,a 130 ± 0.3B,d 27.8 ± 0.2C,de

60 14.1 ± 0.2A,c 4.9 ± 0.1A,a 33.1 ± 0.3A,ab 0.3 ± 0.1A,a 110 ± 0.4C,d 55.4 ± 0.3A,c

Note: PV = peroxide value, p-AV = p-anisidine value, FFA free = fatty acid, IV = iodine value, TEPC= total extracted phenolic content, GalE = gallic acid equivalents, A = 1% alginate,A′= 0.67% alginate, O = canola oil, Q = quercetin, vitamin E = VE, and P = High methoxyl pectin. A–CValues of the same analysis for the same oil sample but different storage periods(Days 0, 30 and 60) with different superscripts are significantly different (p b 0.05). a–fValues of the same analysis and on the same storage day but for different oil samples with differentsuperscripts are significantly different (p b 0.05).

845W. Wang et al. / Food Research International 54 (2013) 837–851

The difference in the chemical structure, solubility, lipophillicity/hydrophicility and emulsifying properties in oils of BHT, vitamin E andquercetin accounted for the detected differences in their correspondingPV, p-AV, FFA and IV results. With hydrophilic groups (e.g. hydroxylgroups), quercetin may migrate and concentrate at the water–oil inter-phase (e.g. the interface between the polysaccharide gel shell and thecore oil),which enables tomore effectively combatwith the penetratingoxygen. During the early stages of storage trials (i.e. Days 0–30), less oilrancidity or degradation of antioxidants that were distributed in hydro-phobic oil occurred.With extended storage (i.e. fromDays 30 to 60), thebreakdown of vitamin E and BHT resulted in the exposure of their phe-nolic units. As a result of quercetin degration, more potent intermediateproducts were possibly generated via ring cleavage. Thus, relativelyhigh readings were detected by Folin–Ciocalteu assay in some caseson Day 60, and in particular, an elevated storage temperature such as38 °C also facilitated increased extractability of phenolics (Sun-Waterhouse et al., 2012). Under harsher storage conditions (38 °C for60 days), more antioxidants were consumed to preserve oil and/orunderwent self-degradation, causing a simultaneous decrease in TEPC.In this study, oil primary oxidation wasmuchmore dominant than sec-ondary oxidation and hydrolytic rancidity. The 1%A and A–P shell for-mulations are acceptable and comparable for preserving quercetin-containing canola oil, with 1%A being slightly better for 60 day storagesand the A–P shell for short storage periods up to 30 days.

3.5. HPLC analysis of phenolics in the encapsulated quercetin-containingcanola oil beads

HPLCanalyses indicate that the shell formulations and storage condi-tions determined the amount of quercetin, flavonoids, non-flavonoidphenolic compounds and some non-phenolic substances (Table 5).The absorption spectra of flavonoids generally consist of two character-istic bands with maxima at 280 and 370 nm (Zvezdanović, Stanojević,Marković, & Cvetković, 2012). Quercetin can be quantified at the370 nmwavelength using rutin as an internal standard,which appearedin the HPLC chromatograms at ~7.2 min and ~3.5 min, respectively.

On Day 0 (Fig. 3A), quercetin (2.16–3.12 μg/g dried bead) and twoflavonoids at 3.3 and 4.1 min were detected in all the 3 types of encap-sulated oil samples. After 30 days at 20 °C (HPLC profiles not shown),no quercetin was identified but instead two flavonoids (at 3.7 and4.1 min) were detected in the 3 types of encapsulated beads. FlavonoidIII (at 4.1 min) was also present in all the Day 0 beads. After 30 days at38 °C (HPLC profiles not shown), neither quercetin nor any other flavo-noid was detected, instead, a non-flavonoid phenolic compound at3.2 min was identified at 280 nm in the 3 types of beads. This phenoliccompound might be a degradation product derived from quercetin.Moreover, non-phenolic substances were found at 4.2, 4.7, 6.8, 8.4and/or 10.2 min in the 3 types of beads. They might originate from theshell materials. The 0.67%A beads had two extra substances at 6.8 and

Table 5Amount of identified compounds in encapsulated quercetin-containing oil beads.

Storage Bead Identified compounds at different wavelengths

370 nm 280 nm

Day 0 0.67% Alginate 7. 2 min, quercetin, 2.17 ± 0.01 μg/g dried bead,4.1 min flavonoid III, 0.82 ± 0.01 μg/g dried bead,3.3 min, flavonoid I, a big shoulder peak.

No detected phenolic and non-phenolic compounds.

1% Alginate 7.2 min quercetin, 3.12 ± 0.02 μg/g dried bead,4.1 min flavonoid III, 0.72 ± 0.02 μg/g dried bead,3.3 min, flavonoid I, a big shoulder peak.

No detected phenolic and non-phenolic compounds.

Alginate–pectin 7.2 min, quercetin, 2.16 ± 0.02 μg/g dried bead,4.1 min flavonoid III, 0.80 ± 0.01 μg/g dried bead,3.3 min, flavonoid I, a big shoulder peak.

4.5 min, a very small phenolic peak;no detected non-phenolic compounds.

Day 30, RT 0.67% Alginate No quercetin detected,3.7 min flavonoid II, 0.79 ± 0.01 μg/g dried bead,4.1 min flavonoid III, 0.09 ± 0.01 μg/g dried bead.

No detected phenolics;an unknown non-phenolic compound at 10.2 min.

1% Alginate No quercetin detected,3.7 min flavonoid II, 0.71 ± 0.01 μg/g dried bead,4.1 min flavonoid III, 0.29 ± 0.02 μg/g dried bead.

No detected phenolic and non-phenolic compounds.

Alginate–pectin No quercetin detected, 3.7 min flavonoid II, 0.99 ± 0.02 μg/g dried bead,4.1 min flavonoid III, 0.06 ± 0.01 μg/g dried bead.

No detected phenolic and non-phenolic compounds.

Day 30, 38 °C 0.67% alginate No quercetin and other flavonoid detected. 3.2 min a phenolic compound, 2.89 ± 0.03 μg/g dried bead;unknown non-phenolic compounds at 4.2, 4.7, 6.8, 8.4 and 10.2 min.

1% Alginate No quercetin and other flavonoid detected. 3.2 min a phenolic compound, 3.34 ± 0.02 μg/g dried bead;unknown non-phenolic compounds at 4.2, 4.7 and 10.2 min.

Alginate–pectin No quercetin and other flavonoid detected. 3.2 min a phenolic compound, 3.13 ± 0.01 μg/g dried bead;unknown non-phenolic compounds at 4.2, 4.7 and 10.2 min.

Day 60, RT 0.67% Alginate No quercetin detected,3.3 min, flavonoid I, a shoulder peak, smaller than Day 0.

3.2 min, a phenolic compound, 1.74 ± 0.01 μg/g dried bead;unknown non-phenolic compounds at 4.7 and 10.2 min.

1% Alginate No quercetin detected,3.3 min, flavonoid I, a shoulder peak, smaller than Day 0.

3.2 min, a phenolic compound, 1.66 ± 0.01 μg/g dried bead;an unknown non-phenolic compound at 10.2 min.

Alginate–pectin No quercetin detected,3.3 min, flavonoid I, a shoulder peak, smaller than Day 0.

3.2 min, a phenolic compound, 3.02 ± 0.02 μg/g dried bead;an unknown non-phenolic compound at 10.2 min.

Day 60, 38 °C 0.67% Alginate No quercetin detected,4.1 min, flavonoid III, 0.61 ± 0.03 μg/g dried bead,3.3 min, flavonoid I, a very small shoulder peak.

Unknown non-phenolic compounds at 4.2, 4.7, 5.5, 5.8, 6.9, 8.4, 8.9,10.2, 12.5 and 13.9 min.

1% Alginate No quercetin detected,4.1 min, flavonoid III, 0.39 ± 0.02 μg/g dried bead,3.3 min, flavonoid I, a very small shoulder peak.

Unknown non-phenolic compounds at 4.2, 4.7, 8.4, 10.2 min.

Alginate-pectin No quercetin detected,4.1 min, flavonoid III, 0.46 ± 0.02 μg/g dried bead,3.3 min, flavonoid I, a very small shoulder peak.

Unknown non-phenolic compounds at 4.2, 4.7, 5.8, 6.8, 8.4, 10.2, 12.5and 13.9 min.

Note: RT = room temperature.

846 W. Wang et al. / Food Research International 54 (2013) 837–851

8.4 min compared with the 1%A and A–P beads. After 60 days at 20 °C,no quercetin was detected, but a flavonoid at 3.3 min and a non-flavonoid phenolic compound at 3.2 min were found in the 3 types ofbeads. Furthermore, only one non-phenolic substance (at 10.2 min)was detected in the 1%A and A–P beads, and two non-phenolic sub-stances (at 4.7 and 10.2 min) were found in the 0.67%A beads. After60 days at 38 °C (Fig. 3B), neither quercetin nor non-flavonoid pheno-lics were found, but two flavonoids (at 3.3 and 4.1 min) were seen inthe 3 types of beads. Ten and eight non-phenolic substances were de-tected in the 0.67%A and A–P beads, respectively, but only four non-phenolic species in the 1%A beads. This result agrees with the storagetrial finding about the better stability of the 1%A beads. The non-phenolic compounds detected at 280 nmmight originate from alginateand/or pectin.

The HPLC results suggest the interplay between storage condi-tions and shell wall formulations (which determined the pathwaysand rates of quercetin decomposition). The hydroxyl group at C-3on the C-ring, adjacent to the 2,3-double bond and the 4-carbonyl,is easily oxidised due to the electron donation of the ketone structureat C-4 (Lin, Wang, Qin, & Bergenståhl, 2007; Rice-Evans, Miller, &Paganga, 1996). Previous studies published in the literature havesuggested the possible phenolic products derived from quercetindecomposition such as 3,4-dihydroxybenzoic (protocatechuic)acid, 1,3,5-trihydroxybenzene (phloroglucinol), 1,3,8-Trihydroxy-9aH,11H-benzofuro[3,2-b]-[1]benzopyran-7,11-dione cyclic ether, 2,5,7,3′,4′-pentahydroxy-3,4-flavandione, 2-(3,4-Dihydroxybenzoyl)-2,4,6-

trihydroxy-3(2H)-benzofuranone, 2-(3′,4′-dihydroxyphenyl)-2,3-dihy-droxyprop-2-en-1-al (Buchner, Krumbein, Rohn, & Kroh, 2006; Leiet al., 2008; Zenkevich et al., 2007).

3.6. FT-IR analysis of encapsulated canola oil beads

FT-IR was used to examine the composition of macromolecules inthe complex bead systems. Fig. 4A shows the FT-IR-spectra of the ingre-dients alginate, HM pectin, quercetin, canola oil and water that wereused to prepare the 3 types of quercetin-containing canola oil beads.All spectra were consistent with literature reports for these ingredients(Che Man & Rohman, 2012; Ismail, Ramli, Hani, & Meon, 2012;Krishnakumar, Prabu, & Sulfikkarali, 2012; Manrique & Lajolo, 2002).

FT-IR spectra (Fig. 4B, C and D) of the freeze dried quercetin-containing oil beads encapsulated by 0.67%A, 1%A and A–P, respectively,were similar and dominated by signals from the core substance (canolaoil) and shell material (alginate and HM pectin). The canola oil signalswere as follows; 3009 cm−1 (C–H stretching vibration of the cis-double bond CH), 2925 cm−1 and 2854 cm−1 (symmetric and asym-metric stretching vibration of the aliphatic CH2 group), 1746 cm−1

(ester carbonyl functional group of the triglycerides), 1465 cm−1 (bend-ing vibrations of the CH2 andCH3 aliphatic groups), 1377 cm−1 (bendingvibrations of CH2 groups), 1238 cm−1 and 1163 cm−1 (stretching vibra-tion of the C–O ester groups) and 723 cm−1 (overlapping of the CH2

rocking vibration and the out-of-plane vibration of cis-disubstituted ole-fins) (Che Man & Rohman, 2012; Muik, Lendl, Molina-Díaz, & Ayora-

847W. Wang et al. / Food Research International 54 (2013) 837–851

Cañada, 2005; Vlachos et al., 2006). The dominant oil signals reflect thehigh mass fraction of canola oil in the encapsulated beads especially inthe freeze dried form. As seen in Fig. 4A, the assignment of the absorptionbands for HM pectin in the beads was difficult, as most of these bandsoverlapped with those of alginate and canola oil that were at higherconcentration than HMpectin. Unambiguous assignment of FT-IR absor-bance signals for quercetin was not possible because of its low concen-tration in these freeze dried oil beads. Reactions between quercetin orits degradation products, and the core oil encapsulated by alginatealone or alginate-pectin matrix during storages are evident for data inTables 3–5 (Sun-Waterhouse et al., 2013). Storage at 20 °C or 38 °C for

A

A′+O

A′+O

A+O+

A+O+

A+P

A+P

Fig. 3. High performance liquid chromatograms (at 280 and 370 nm) for 0.67% alginate(A+ P+ O+Q) encapsulated beads on A) Day 0, B) Day 60 at 38 °C.

30 and 60 days did not cause major changes in the FT-IR spectra of allthe 0.67%A, 1%A and A–P beads, reflecting the chemical stability of theencapsulated beads.Where the intensity of oil-related signals decreased,there seemed to be an increase of water-related signals i.e. the bands at3300 and 1635 cm−1 due to asymmetric O–H stretching and H–O–Hbending modes, respectively (Thygesen, Løkke, Micklander, & Engelsen,2003). The 3009 cm−1 signal is associated with the unsaturationdegree of canola oil (Muik et al., 2005), slightly decreased in somecases (e.g. the 0.67%A and A–P bead after 60 days at 38 °C). The intensityof the 723 cm−1 band (overlapping of the CH2 rocking vibration and theout-of-plane vibration of cis-disubstituted olefins) generally remained

+Q, Day 0,280 nm

+Q, Day 0,370 nm

Q, Day 0,280 nm

Q, Day 0,370 nm

+O+Q, Day 0, 280 nm

+O+Q, Day 0, 370 nm

(A′+O+Q), 1% alginate (A + O+ Q) and alginate + high methoxyl (HM) pectin

B

A'+O+Q, Day 60, 38°C, 280 nm

A'+O+Q, Day 60, 38°C, 370 nm

A+O+Q, Day 60, 38°C, 280 nm

A+O+Q, Day 60, 38°C, 370 nm

A+P+O+Q, Day 60, 38°C, 280 nm

A+P+O+Q, Day 60, 38°C, 370 nm

Fig. 3 (continued).

848 W. Wang et al. / Food Research International 54 (2013) 837–851

strong for the 1%A and A–P beads (Fig. 3C and D), but decrease for the0.67%Abeads (Fig. 4B) after storage. These results agreewith thefindingsin storage trials that the 1%A and A–P beads offered better oil protection.

In general, polysaccharide polymer gels create a cross-linked hydro-philic network matrix with a porous structure, formed via chemicalinteractions (e.g. covalent bonds) or physical interactions (e.g. non-covalent hydrogen bonding, hydrophobic, and ionic interactions)(Whistler & BeMiller, 1997). In the network structures, interactions be-tween molecules of the same type are favoured over those betweenmolecules of different types forming “permanent” junction zones or

“temporary” entanglements in the network (Choi & Yoo, 2008). HMpectins form rapid-set gels mainly by hydrophobic interactions and hy-drogen bonds (Oakenfull & Scott, 1984), which may facilitate a rapidseparation (also a reduced contact) between canola oil and oxygen inair, causing decreased oil deterioration. With increasing temperature,the increase in entropy reduces the hydration of pectin chains, and hy-drophobic interactions are strengthened and become the most impor-tant contribution to the macromolecular interactions (Oakenfull &Fenwick, 1977). Thus, the addition of HM pectin to alginate shellmight enhance the stability of the shell and core oil during prolonged

Wavenumber (cm-1)1000150020002500300035004000

Abs

orba

nce

(arb

itrar

y un

its)

Canola oil

Quercetin dihydrate

HM pectin

Sodium alginate

Water

Wavenumber (cm-1)1000150020002500300035004000

Abs

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nce

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its)

A'+O+Q day 0

A'+O+Q day 30 RT

A'+O+Q day 30 38ºC

A'+O+Q day 60 RT

A'+O+Q day 60 38ºC

Wavenumber (cm-1)1000150020002500300035004000

Abs

orba

nce

(arb

itrar

y un

its)

A+O+Q day 0

A+O+Q day 30 RT

A+O+Q day 30 38ºC

A+O+Q day 60 RT

A+O+Q day 60 38ºC

Wavenumber (cm-1)1000150020002500300035004000

Abs

orba

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(arb

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its)

A+P+O+Q day 0

A+P+O+Q day 30 RT

A+P+O+Q day 30 38ºC

A+P+O+Q day 60 RT

A+P+O+Q day 60 38ºC

A, ingredients B, 0.67% alginate

D, alginate-pectinC, 1% alginate

Fig. 4. Normalised FT-IR absorbance spectra for A) ingredients, and freeze dried encapsulated quercetin–canola oil beads before and after a storage at room temperature (RT) or 38 °C:B) 0.67% alginate alone (A′ + O + Q), C) 1% alginate alone (A + O + Q), D) alginate + high methoxyl (HM) pectin (A + P + O + Q). The spectra have been normalised and offsetvertically for clarity.

849W. Wang et al. / Food Research International 54 (2013) 837–851

storage especially at higher temperature (e.g. 38 °C) (Endress &Rentschler, 1999). However, a remarkable difference in the behaviourof HM pectin as a function of storage temperature is not expected inthis study due to the small temperature difference between 20 °C and38 °C. In comparison, the alginate alone gels where hydrogen bonding

or electrostatic interactions are the only significant interactions forstabilising the polymer network (Andresen & Smidsrod, 1977), a mo-notonous decrease of the elastic modulus with increasing temperatureis expected. Hence, the alginate alone oil beads may be less robustthan A–P beads as a function of temperature.

850 W. Wang et al. / Food Research International 54 (2013) 837–851

4. Conclusions

Co-extrusion encapsulation using alginate and alginate-HM pectinshell materials is a feasible approach for encapsulating unsaturated ca-nola oil. Encapsulation conditions influenced the characteristics of theoil beads, with the core and shell flow rates being the critical to realisingstable spherical oil beads. Bead size, core oil stability and retained phe-nolic content vary with the shell wall formulations. Core oil fortifiedwith quercetin is more effective than fortification with vitamin E orBHT for suppressing oil oxidation at 38 °C. FT-IR studies confirmedthe chemical stability of the three types of quercetin-containing encap-sulated beads during storage. HPLC analyses revealed that the degrada-tion pathway of quercetin depended on the shell formulation andstorage conditions.

Both 1% alginate and alginate–HM pectin shell formulations are ac-ceptable and comparable for preserving quercetin-fortifying canola oil.Considering the nutritional value introduced by the pectin addition,the alginate–pectin shell represents a good option for encapsulating un-saturated oil for food ingredient application. The quercetin-containingcanola oil beads encapsulated with alginate-based polysaccharideshell, which can be consumed directly or used as bioactive ingredients,simultaneously deliver the goodness of unsaturated oil, quercetin anti-oxidant and fibre polysaccharides to consumers.

Acknowledgments

We acknowledge research funding from the University of Aucklandand Plant & Food Research.

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