Food Chemistry 158 (2014) 497–503

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Food Chemistry

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Analytical Methods

The determination of botanical origin of honeys based on enantiomerdistribution of chiral volatile organic compounds

http://dx.doi.org/10.1016/j.foodchem.2014.02.1290308-8146/� 2014 Elsevier Ltd. All rights reserved.

⇑ Corresponding author. Tel.: +421 2 59325283; fax: +421 2 52926043.E-mail address: ivan.spanik@stuba.sk (I. Špánik).

Ivan Špánik a,⇑, Alexandra Pažitná a, Peter Šiška b, Peter Szolcsányi ba Slovak University of Technology, Faculty of Chemical Technology, Institute of Analytical Chemistry, Radlinského 9, SK-812 37 Bratislava, Slovak Republicb Slovak University of Technology, Faculty of Chemical Technology, Department of Organic Chemistry, Radlinského 9, SK-812 37 Bratislava, Slovak Republic

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

Article history:Received 1 February 2013Received in revised form 18 September2013Accepted 23 February 2014Available online 6 March 2014

Keywords:HoneyBotanical originMultidimensional gas chromatographySeparation of enantiomersSolid phase microextraction (SPME)Chiral volatile organic compounds (VOC’s)

The enantiomer ratios of chiral volatile organic compounds in rapeseed, chestnut, orange, acacia,sunflower and linden honeys were determined by multi-dimensional gas chromatography using solidphase microextraction (SPME) as a sample pre-treatment procedure. Linalool oxides, linalooland hotrienol were present at the highest concentration levels, while significantly lower amounts ofa-terpineol, 4-terpineol and all isomers of lilac aldehydes were found in all studied samples. On the otherhand, enantiomer distribution of some chiral organic compounds in honey depends on their botanicalorigin. The significant differences in enantiomer ratio of linalool were observed for rapeseed honey thatallows us to distinguish this type of honey from the other ones. The enantiomer ratios of lilac aldehydeswere useful for distinguishing of orange and acacia honey from other studied monofloral honeys.Similarly, different enantiomer ratio of 4-terpineol was found for sunflower honeys.

� 2014 Elsevier Ltd. All rights reserved.

1. Introduction

Honey is probably one of the most important natural productthat is widely used not only in food industry, but also in medicinedue to its antibacterial properties, in cosmetics as a part of hydrat-ing or cold lotion or as a raw material for production of honeywine. One can distinguish between monofloral and polyfloralhoney. Monofloral honey is usually more appreciated, since it pri-marily originates from the nectar of one type of flower. Monofloralhoneys differ in taste, flavor as well as in colour depending onproperties of primary nectar sources.

Up to these days, various analytical methods have been usedfor determination of botanical origin of monofloral honeys(Kaskoniene & Venskutonis, 2010) e.g. analysis of pollen (melissop-alynology) (Sabo, Potocnjak, Banjari, & Petrovic, 2011) or monitor-ing of various analytical parameters, like sugar content (Kukurová,Karovičová, Kohajdová, & Bílikova, 2008), profile of phenoliccompounds and flavonoids (Escriche, Kadar, Juan-Borras, &Domenech, 2011), conductivity, mineral content, pH and colour(Vanhanen, Emmertz, & Savage, 2011), stable isotopes (Rossmann,2001) etc., but often a correct determination of honey originrequires a detailed knowledge of its physical properties and

chemical composition. In the past decade, it was shown that com-position of volatile organic compounds (VOCs) has to be consideredas a perspective tools in the assessment of honey origin. It is notsurprising, since volatile fraction of honey contains more than150 organic compounds (Bentivenga, D’Auria, Fedeli, Mauriello &Racioppi, 2004; Castro-Vazquez, Perez-Coello, & Cabezudo, 2003;Špánik et al., 2013). The composition of volatile organic com-pounds in honey mostly depends on the botanical origin of flowersfrom which honey was made. Indeed, many papers publishedwithin last 10 years deal with characterization of VOC profile ofhoneys with different botanical origin with special attention tothe identification of their possible markers. For example, in 2010Aliferis, Tarantilis, Harizanis, and Alissandrakis (2010) have identi-fied lilac aldehyde and the 1-p-menthen-9-al isomers, limonene,methyl anthranilate, phenyl acetaldehyde, 1-phenyl-2,3-butanedi-one, 3-hydroxy-4-phenyl-2-butanone, 3-hydroxy-1-phenyl-2-butanone, and 3-hydroxy-4-phenyl-3-buten-2-one as possiblemarkers for classification of citrus and thyme honeys. Similarly,in 2001 Radovic et al. (2001) studied possibility to differentiateacacia, chestnut, eucalyptus, heather, lime, lavender, rapeseed,rosemary, and sunflower honeys based on the presence or absenceof certain VOCs. The characteristic VOC profiles were also usedto discriminate between Corsican and non-Corsican honeys(Stanimírova et al., 2010).

Considering the fact, that VOCs in honey is a complex mixture ofdifferent compounds containing various chemical functional

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498 I. Špánik et al. / Food Chemistry 158 (2014) 497–503

groups with relatively low molecular weight that are present atlow concentration levels, their extraction requires utilisation ofeffective sample treatment methods. There are several ways howto extract VOCs from honey in order to reach required sensitivity,recovery and reliability. The most frequently used is solid phasemicroextraction. This solventless technique can be easily auto-mated, but requires a time consuming optimization of workingconditions like the type of SPME fibres, sorption temperature, sorp-tion time and desorption temperature. Currently, SPME is mostlyused for the characterization of honey VOC profiles (Kaskoniene,Venskutonis, & Ceksteryte, 2008; Plutowska, Chmiel, Dymerski, &Wardencki, 2011; Soria, Martinez-Castro, & Sanz, 2003; Soria, Sanz,& Martinez-Castro, 2009). Another extraction method is liquid–liquid extraction into organic solvent which was widely usedbefore invention of SPME. The major advantage of this method isits simplicity and ability to extract thermolabile compounds thatdo not withstand an extensive heating. The selectivity of this meth-od is mostly influenced by selection of proper organic solvent thatallows tuning of extraction conditions only for particular com-pounds with similar chemical properties (Rowland, Blackman,D’Arcy, & Rintoul, 1995; Wilkins, Lu, & Tan, 1993). Some papersalso report simultaneous steam distillation extraction method(Bicchi, Belliardo, & Frattini, 1983), static headspace (Cuevas-Glory,Pino, Santiago, & Sauri-Duch, 2007) or purge and trap (Soria,Martinez-Castro, & Sanz, 2009).

It was demonstrated that many organic compounds present inhoney are chiral, thus can exist as pair of enantiomers. These chiralcompounds are mostly synthesized in plants by enzymatic reac-tions. The determination of enantiomer composition of chiral com-pounds becomes more utilised for evaluation of adulteration, andmanipulation with non-food commodities or to differentiate be-tween natural and synthetic compounds. This method is basedon many observations, one of them being that chiral compoundsoccur in the nature as pure enantiomers or mixture of enantiomerswith specific ratio. Any changes in these ratios indicate illicitmanipulation with products or addition of synthetically producedchemicals. In food industry, this approach has been firstly usedfor the identification of adulteration of orange juices by additionof synthetic amino acids, which contain also D-enantiomers, usu-ally not present in the nature (Ooghe, Kasteleyn, Temmerman, &Sandra, 1984). Since 1984, only a few papers have been orientedon the utilisation of enantiomer composition as authenticationmarkers. Noticeable changes start since 2002, when the use ofenantiomer distribution of flavours and fragrances as possibleauthentication markers of geographical origin (del Castillo,Caballero, Blanch, & Herraiz, 2002; del Castillo, Caja, Blanch, &Herraiz, 2003), used declared technological procedure (del Castillo,Flores, Herraiz, & Blanch, 2003a) or adulteration by synthetic aro-mas (del Castillo, Flores, Herraiz, & Blanch, 2003b) has been stud-ied. Pažitná et al., in her latest study from 2012 have shown thatone dimensional gas chromatography with chiral stationary phaseshas a limited ability to determine the correct enantiomer distribu-tion in complicated food matrices. In such cases, even if chiralstationary phase shows an excellent efficiency towards enantiomerseparation, resolved enantiomers often coelute with anothernon-chiral or already separated enantiomer of another organiccompound. This lack of separation efficiency can be overcome bytwo dimensional approach, in which two independent GC ovensequipped with proper switching system (e.g. heart-cut) andcolumn setup (first column with achiral stationary phase andsecond column with chiral stationary phase). Such GC system ismore suitable for the determination of enantiomer compositionof organic compounds present in complex matrices (Pažitná,Janáčova, & Špánik, 2012).

The aim of this work was to determine enantiomer ratio ofselected volatile organic compounds in acacia, rapeseed, sunflower,

linden, chestnut and orange honeys and to investigate the possibil-ity to use them for evaluation of their botanical origin.

2. Materials and methods

2.1. Samples

A monofloral honey samples were obtained at local marketsfrom local beekeepers in various European countries. Those were(i) acacia honey (Robinia pseudoacacia) – totally 12 samples (3 fromSlovakia, 2 from Czech Republic and Romania and 1 sample fromGermany, Serbia, Georgia, Poland and Moldova), (ii) rapeseedhoney (Brassica napus) – 3 samples from Slovakia, (iii) sunflowerhoney (Helianthus annuus) – 2 samples from Slovakia and 1 samplefrom Ukraine, (iv) linden honey (Tilia cordata) – totally 7 samples(2 from Slovakia, 1 sample from Czech Republic, Romania, Hungaryand Moldova, (v) chestnut honey (Aesculus hippocastanum) – 2samples from Italy and finally (vi) orange honey (Citrus aurantium)– 2 samples from Greece and 1 sample from both Italy and France.All samples underwent analysis of their basic physicochemicalparameters such as sugar content, pH, conductivity, acidity, andwater content.

2.2. Sample treatment

VOCs from honey samples were extracted by SPME procedureusing Agilent autosampler GC 80. A 5 g of honey together with a0.5 g of NaCl were dissolved in DI water in a 20 ml clear glass vialsand the solution was stirred with a PTFE-coated magnetic stir barat 450 rpm. Vials were sealed with hole-caps and PTFE/siliconesepta. A stirred sample solution was heated at 60 �C for 30 minin order to establish equilibrium between liquid and vapourphases. VOCs were extracted by SPME fibre for autosampler coatedwith PDMS/CAR/DVB (50/30 lm thickness) obtained from Supelco(Bellefonte, PA, USA). The fibre was conditioned prior use by heat-ing in the injection port of GC under the conditions recommendedby the manufacturer. The adsorption of VOCs from honey sampleson SPME fibre took 30 min at 60 �C while solution was stirred at450 rpm. Desorption was performed in GC injector in splitlessmode at 220 �C for 2 min.

2.3. Gas chromatography

Gas chromatograph Agilent 7890A connected to Agilent 5975CMS detector was used in GC–MS experiments. The sample wasintroduced into GC via split/splitless injector heated at 250 �Cworking in splitless mode by Agilent SPME autosampler GC 80.Helium with purity 99.995% was used as a carrier gas with a flowrate 1 ml min�1. A 30 m DB-FFAP column (Agilent J & W Column,Agilent Technologies, USA) with i.d. 0.25 mm and film thickness0.25 lm was used in GC–MS experiments. A temperature programstarted at 60 �C, after then temperature increased with a gradient2 �C/min to 150 �C followed by 10 �C/min to 250 �C.

Enantiomer separations were performed in two dimensional GCsystem containing two independent GC ovens. In this approach theVOCs extracted from honeys were firstly separated in GC oven atachiral stationary phase. When chiral compound eluted from thefirst column, a switching system redirected this compound to thesecond GC oven with chiral stationary phase. The first gaschromatograph, identical with this used in GC–MS experiments,was connected via Dean’s microfluidic switching system (AgilentTechnologies, USA) to the second gas chromatograph HewlettPackard 5890, which was equipped with FID detector. Helium withpurity 99.995% was used as a carrier gas with a flow rate in theDB-FFAP column 1 ml min�1. The carrier gas flow was switched

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in Dean’s microfluidic device either to the restrictor (length 8.8 m,i.d. 0.18 mm) that ends in MS detector, or, if chiral compound haseluted from DB-FFAP column, either to 20 m capillary column with0.25 mm i.d. coated with permethyl-b-cyclodextrin anchored tosilicone polymer – ChirasilDex (CHROMPACK International, TheNetherlands), or, in case of lilac aldehydes, to 25 m capillary col-umn with 0.25 mm i.d. coated with 6-tert.butyldimethylsilyl-2,3-dimethyl-b-cyclodextrine dissolved in polysiloxane (MEGA, Italy)inserted in the second GC oven. The VOC separation in the firstGC oven was performed under the same conditions as GC–MSexperiments. Chiral separations of selected compounds were per-formed isothermally at 50 �C (linalool), 80 �C (lilac aldehydes) or75 �C (all other chiral compounds).

2.4. Chemicals

Anhydrous NaCl used in sample treatment procedure was ob-tained from Mikrochem (Pezinok, Slovakia). DI water was preparedin laboratory using NANOpure device from Wilkem Werner GmbH.The racemic mixtures of linalool, 4-terpineol, a-terpineol, furanoidcis-(2R,5S; 2S,5R) and trans-(2R,5R; 2S,5S) linalool oxides, as well asenantiomerically pure (R)-linalool, (S)-4-Terpineol and (S)-a-Terpineol were obtained from Sigma Aldrich (St. Louis, MO, USA).The racemates of lilac aldehydes isomer A (2S,20S,50S; 2R,20R,50R),isomer B (2R,20S,50S; 2S,20R,50R), isomer C (2R,20S,50R; 2S,20R,50S)and isomer D (2S,20S,50R; 2R,20R,50S) were synthetically preparedaccording to procedures published by Kreck and Mosandl (2003).The racemate of hotrienol was synthetically prepared from linalolacetate by described procedure (Yuasa & Kato, 2003). The enantio-merically pure cis-(2S,5R)-linalool oxide and trans-(2R,5R)-linalooloxide were prepared form (R)-linalool according to procedure de-scribed by Urones et al. (1995). On the other hand, (2R,20R,50R)-lilacaldehyde A, (2S,20R,50R)-lilac aldehyde B, (2R,20S,50R)-lilac aldehydeC, and (2S,20S,50R)-lilac aldehyde D were prepared from (R)-linaloolvia adapted 2-step sequence (allylic oxidation followed by cyclisa-tive Michael addition) according to known synthetic protocols(Kreck, Püschel, Wüst, & Mosandl, 2003; Matich et al., 2003). Thestudied VOC’s were identified by comparison of measuredretention times and mass spectra with those obtained for racemicmixtures of standards injected under identical conditions. Theelution order of linalool, lilac aldehydes and linalool oxides was

Fig. 1. Comparison of extraction efficiencies for selec

determined by injection of racemic mixture enriched by the corre-sponding pure enantiomer. The obtained elution order was alsocompared to those published in literature for those of othercompounds on permethylated-b-CD (Baigrie, Chisholm, &Mottram, 1996; Demyttenaere & Willemen, 1998; Schurig &Nowotny, 1988) or on 6-tert-butyldimethylsilyl-2,3-dimethyl-b-cyclodextrine (Kreck & Mosandl, 2003), except of hotrienol, whichaccording to our best knowledge, is not available in the literature.

3. Results and discussion

Linalool, hotrienol, lilac aldehydes, 4-terpineol, a-terpineol,cis- and trans-linalool oxides have been reported in previous pa-pers as major constituents of VOCs found in honeys independentlyof their botanical origin (Castro-Vazquez et al., 2003; Soria et al.,2003; Stanimírova et al., 2010). Thus, the first part of our workwas focused at optimization of SPME procedure to reach efficientextraction of those VOC’s from honey samples. The optimizationwas performed on one sample of acacia honey. During optimiza-tion of SPME working conditions, type of SPME coatings (PA, PDMS,PDMS/DVB and CAR/PDMS/DVB), sorption temperature and timehave been studied. Fig. 1 illustrates a comparison of measured peakareas for given VOC reached on particular SPME coatings. PDMS/DVB showed the best results in terms of number of extracted VOCsas well as their quantity. This observation is in agreement withpreviously published results (Soria et al., 2003; Čajka, Hajšlová,Cochran, Holadová, & Klimánková, 2007). The number of extractedcompounds as well as their peak areas has increased with increas-ing sorption temperature from 30 to 90 �C showing a small maxi-mum at 60 �C. Fig. 2 illustrates a comparison of measured peakareas for studied chiral compounds obtained at various sorptiontemperatures. However, in the follow-up experiments the optimalsorption temperature of 60 �C was selected to prevent a possibledecomposition of thermolabile compounds. Similarly, thoseparameters also increased with increasing sorption time from 10to 30 min. After then only small changes in measured peak areaswith increasing sorption time were observed. Slow temperaturegradient (2 �C/min) was chosen to reach the best resolution ofVOC’s on achiral stationary phase. Thus, the optimization ofseparation conditions was mostly focused on selection of properstationary phase. The most frequently used stationary phase for

ted VOC’s obtained on different SPME coatings.

Fig. 2. Comparison of measured peak areas for studied chiral compounds obtained at various sorption temperatures.

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separation of VOC’s from honey samples includes non-polar 5%fenyl – 95% dimethyl polysiloxane (HP-5, 30 m, i.d. 0.25 mm, filmthickness 0.25 lm) or polar modified polyethylene glycol station-ary phase (DB- FFAP). In our work, both stationary phases wereused. Both columns have showed similar separation efficiencies,however only modified polyethylene glycol stationary allowed usto separate B and C isomers of lilac aldehydes. Fig. 3 illustratesthe chromatogram obtained for acacia honey on DB-FFAP columnunder optimised working conditions. The chromatograms of otherstudied honeys are presented in Supplementary materials(Figs. 1S–5S). Totally, more than 270 compounds were detectedthat belong to various chemical classes, e.g. aliphatic and branchedalkanes, alcohols, aldehydes, ketones, carboxylic acids and theirmethyl and/or ethyl esters etc. Linalool oxides, linalool and hotrie-nol were present at the highest concentration levels, while signif-icantly lower amounts of a-terpineol, 4-terpineol and all isomersof lilac aldehydes were found. Above mentioned volatile organic

Fig. 3. The chromatogram obtained for acacia honey on DB-FFAP column under optimise4-linalool 5-lilac aldehyde B, 6-lilac aldehyde C, 7-lilac aldehyde D, 8–4-terpineol, 9-hot

compounds were present in all studied honey samples, which isin agreement with previously reported results (Špánik et al.,2013). The relative content of selected VOC’s in studied samplesis shown in Fig. 4.

The enantiomer ratios of each compound were determinedusing multidimensional gas chromatography in separate GC runs.Fig. 5 illustrates chromatograms of separated enantiomers of race-mic mixtures of studied VOC’s and obtained enantiomers ratios areshown in Table 1. It can be seen that obtained enantiomer ratiosvary within wider range (generally up to 10%). It can be concludedthat differences in distribution of enantiomers of studied chiralVOC’s in some cases depends on botanical origin of honey.(2R,5S)-cis-linalool oxide was predominant enantiomer in all stud-ied honeys, however, only in linden honey its content exceed 80%with respect to its (2S,5R) enantiomer. Almost racemic mixture oftrans-linalool oxide was found in rapeseed, orange, acacia, andlinden honeys. In sunflower honey, slight predomination of

d working conditions. 1-trans-Linalool oxide, 2-cis linalool oxide, 3-lilac aldehyde A,rienol, 10-alpha-terpineol.

Fig. 4. The relative content of selected VOC’s in honey samples based on their botanical origin.

Fig. 5. Enantiomer separation of racemic mixtures of studied VOC’s.

Table 1The percentual content of first eluted enantiomer in studied honey samples.

Compound name Rapeseed Chestnut Sunflower Orange Acacia Linden

(2R, 5S)-cis-Linalool oxide 63–74 42–48 63–75 67–78 60–69 80–93(2R,5R)-trans-Linalol oxide 45–58 19–22 66–72 51–62 44–62 43–58(R)-Linalool 51–66 12–18 27–48 20–33 19–30 16–37Hotrienol 9–15 68–75 6–8 15–18 70–92 7–15(S)-4-Terpineol 48–57 28–31 58–84 25–44 41–65 24–30(S)-a-Terpineol 52–53 62–64 57–61 62–72 52–62 77–86(2S,20S,50S)-Lilac aldehyde A 68–72 53–59 60–64 85–91 61–70 78–87(2R,20S,50S)-Lilac aldehyde B 25–32 32–37 27–34 7–10 4–15 15–29(2R,20S,50R)-Lilac aldehyde C 41–43 38–40 40–45 10–16 27–45 22–29(2S,20S,50R)-Lilac aldehyde D 37–45 77–82 31–35 10–25 24–45 15–24

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(2R,5R)-trans-linalool oxide over its second enantiomer was ob-served. Only chestnut honeys from Italy were characteristic withpredomination of (2S,5S)-enantiomer of trans-linolool oxide. Thuswe conclude that the enantiomer ratio of trans-linalool oxide couldbe used as a marker for sunflower and chestnut honeys. The pres-ence of both enantiomers of furanoid linalool oxides in the specificratio was explained by Luan, Mosandl, Gubesch, and Wüst (2006)by two different pathways that are used to generate furanoid linal-ool oxides in plants. He also observed racemization of furanoid lin-alool oxides from grapes during ageing of wine. Similarly, anotherbiochemical processes connected to the conversion of nectar tohoney in bee hives can contribute to racemisation or changes inenantiomer ratios of studied compounds. (R)-linalool was predom-inant only in rapeseed honey, while in all other studied sampleshigher content of the (S)-enantiomer was observed. The highestenantiomeric purity of (S)-linalool was found in chestnut honeys.This is not surprising; since it was proven that some bee speciesproduces pure (S)-enantiomer as a male attractant (Borg-Karlsonet al., 2003). Acacia and chestnut honeys were characteristic byhigher content of the first eluted enantiomer of hotrienol, whilein other honeys the second eluted enantiomer was present inrelatively high enantiomeric purity. It is supposed that this surpris-ingly high enantiomeric purity of hotrienol is caused by catalyticeffect of specific enzyme (Luan et al., 2006). The distribution ofenantiomers of 4-terpineol was useful for the determination ofsunflower honey origin, where the (S)-enantiomer is predominant.Chestnut and linden honeys were characteristic by presence of sig-nificantly higher amount of (R)-4-terpineol. In rapeseed, acacia andorange honeys almost racemic mixtures were observed. Lilac alde-hydes can be biosynthetized in plants from linalool using specificenantioselective enzymes (Doetterl et al., 2006). Since some plantsproduce (R)-linalool and bee produce (S)-linalool, different enan-tiomeric ratios of lilac aldehydes in honeys could be expected.The first eluted enantiomer of (2S,20S,50S)-lilac aldehyde A waspredominant in all studied samples. The highest content of thisparticular enantiomer was observed in orange and linden honeysthat were significantly different in comparison to honeys of otherbotanical origin. On the contrary, the second eluted (2S,20R,50R)-lilac aldehyde B and (2S,20R,50S)-lilac aldehyde C were predominantin all studied samples. These enantiomers can be used to distin-guish between orange and acacia honeys, where they are presentin highest enantiomeric purity. The first eluted (2S,20S,50R)-lilacaldehyde D can be used as a marker of chestnut honey, since itwas predominant only in this type of honey. On the other hand,the second eluted enantiomer having the (2R,20R,50S)-configurationwith highest enantiomeric purity was found in orange and lindenhoney.

4. Conclusions

In this study, GC–MS was used to characterise VOC fractions ofacacia, sunflower, linden, rapeseed, chestnut and orange honeysform various European countries. Totally, more than 270compounds were detected in studied honeys that belong to variouschemical classes. The most abundant were aliphatic and branchedalkanes, alcohols, carboxylic acids and their methyl and/or ethylesters, while many of them were chiral. Among those, linalool oxi-des, linalool and hotrienol were present at the highest concentra-tion levels. The other chiral compounds (a-terpineol, 4-terpineoland all isomers of lilac aldehydes) were present in significantlylower amounts. The enantiomeric ratios of chiral volatile organiccompounds were determined by multi-dimensional gaschromatography. For honeys of the botanical origin, the obtainedenantiomeric ratios varied in range up to 10%. Importantly, thesedifferences in distribution of enantiomers of studied chiral VOC’s

were dependent on the botanical origin of honey. Thus, our obser-vation holds a potential to be used in a distinguishing of honey’sbotanical origin. However, for the practical purposes, it isrecommended to determine the honey’s origin on the basis of theenantiomeric ratios obtained for larger pool of chiral VOC’s. Ithas to be noted, that it was not possible to determine geographicalorigin of unifloral honeys, since observed enantiomeric ratios wereindependent of country of origin.

Acknowledgements

This work was supported by VEGA grant No. 1/0972/12 andScience and Technology Assistance Agency under contract No.APVV-0428-12. Alexandra Pažitná and Peter Šiška would like tothank Slovak University of Technology for grant supporting youngresearchers.

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.2014.02.129.

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The determination of botanical origin of honeys based on enantiomer distribution of chiral volatile organic compounds1 Introduction2 Materials and methods2.1 Samples2.2 Sample treatment2.3 Gas chromatography2.4 Chemicals

3 Results and discussion4 ConclusionsAcknowledgementsAppendix A Supplementary dataReferences