Storage of extra virgin olive oil and its effect on the biological activity and concentration of oleocanthal

Food Research International 50 (2013) 597–602

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Food Research International j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / f o o d r e s

Storage of extra virgin olive oil and its effect on the biological activity and concentration of oleocanthal Sara Cicerale a, Xavier A. Conlan b, Neil W. Barnett c, Russell S.J. Keast a,⁎ a b c

School of Exercise and Nutrition Sciences, Deakin University, Victoria, Australia Institute for Technology, Research and Innovation, Deakin University, Victoria, Australia School of Life and Environmental Sciences, Deakin University, Victoria, Australia

a r t i c l e

i n f o

Article history: Received 6 December 2010 Received in revised form 21 March 2011 Accepted 23 March 2011 Keywords: Oleocanthal Extra virgin olive oil Phenolics Storage HPLC Taste bioassay

a b s t r a c t The olive oil phenolic, oleocanthal has recently received attention regarding its anti-inflammatory capacity and the thought that it is partially responsible for the beneficial health effects of the Mediterranean diet. Extra virgin olive oil (EVOO) containing oleocanthal is often consumed after storage for a substantial amount of time and for oleocanthal to indeed provide health benefits, it has to be present in a substantial quantity throughout the shelflife of EVOO. Therefore, the aim of the current study was to investigate if natural light and/or oxygen (O2) exposure (via atmospheric air) combined with extended storage, as would occur on a common domestic basis, affected the concentration of oleocanthal in EVOO. One EVOO containing 90 mg/kg oleocanthal was stored for 10 months with and without exposure to light and or O2. Oleocanthal concentrations were quantified using high performance liquid chromatography–mass spectrometry (HPLC–MS) and its biological activity determined with a taste bioassay measuring the intensity of oropharyngeal irritation. A significant difference in oleocanthal concentration was observed amongst the different storage treatments (p = 0.05). Oleocanthal concentration degraded to a maximum of 37% (90 ± 13 mg/kg to 56± 9 mg/kg) after 10 months exposure to both light and O2. Limiting light and O2 over 10 months resulted in a 15% decrease in oleocanthal (90 ± 13 mg/kg to 76 ± 9 mg/kg). Oleocanthal biological activity mirrored the results of oleocanthal concentration (r= 0.8, p b 0.05). Overall, the findings support the role of oleocanthal as a potential health promoting compound in EVOO as significant concentrations remain in EVOO after exposure to light, oxygen and over time. © 2011 Elsevier Ltd. All rights reserved.

1. Introduction Extra virgin olive oil (EVOO) is a major lipid component of the Mediterranean diet (Willett et al., 1995), and there is a wealth of epidemiological evidence demonstrating that Mediterranean populations have reduced risk for certain chronic diseases (such as atherosclerosis, cardiovascular disease (CVD), and particular types of cancer), and extended life expectancy compared with other geographic populations (La Vecchia, 2004; Matalas, Zampelas, Stavrinos, & Wolinsky, 2001; Panagiotakos et al., 2004). These health benefits have been partially attributed to a high dietary consumption of EVOO (characteristically 25–50 ml per day) (Corona, Spencer, & Dessi, 2009). Historically, the healthful properties of EVOO were attributed to its high monounsaturated fatty acid (MUFA) content (Tripoli et al., 2005). However, recent investigations have turned to olive oil phenolics as several seed oils, high in MUFA but devoid in phenolics, have been demonstrated to be ineffective in lowering the risk of ⁎ Corresponding author at: School of Exercise and Nutrition Sciences, Deakin University, 221 Burwood Highway, Burwood, Victoria 3125, Australia. Tel.: +61 3 9244 6944; fax: +61 3 9244 6017. E-mail address: [email protected] (R.S.J. Keast). 0963-9969/$ – see front matter © 2011 Elsevier Ltd. All rights reserved. doi:10.1016/j.foodres.2011.03.046

developing certain chronic diseases (e.g. CVD) (Aguilera et al., 2004; Harper, Edwards, & Jacobson, 2006). Furthermore, studies (including human, animal, in vivo and in vitro) have shown that olive oil phenolics have beneficial effects on certain physiological parameters, such as plasma lipoproteins, oxidative damage, inflammatory markers, platelet and cellular function, antimicrobial activity, and bone health. For reviews, see articles by Cicerale et al. (Cicerale, Conlan, Sinclair, & Keast, 2009; Cicerale, Lucas, & Keast, 2010). An olive oil phenolic of current interest due to its suggested health benefiting properties is oleocanthal. This compound possesses an antiinflammatory capacity due to its dose-dependent ability to inhibit cyclooxygenase (COX) enzymes involved in the prostaglandin biosynthesis (inflammatory) pathway (Beauchamp et al., 2005). Therefore, it has been hypothesised that chronic long-term ingestion of low doses of oleocanthal may be responsible, in part, for the lowered incidence of heart disease, certain cancers, and other degenerative diseases associated with the Mediterranean diet (Beauchamp et al., 2005). In support of health benefits associated with oleocanthal, in vitro studies have demonstrated effective inhibition of the proliferation, migration, and invasion of human breast and prostate cancer lines (Elnagar, Sylvester, & El Sayed, 2011). Furthermore, oleocanthal has been shown to possess therapeutic activities that would be beneficial in the

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treatment of Alzheimer's disease (Li et al., 2009; Pitt et al., 2009), and this supports research demonstrating a 40% decrease in Alzheimer's in populations consuming a Mediterranean based diet (Scarmeas et al., 2009). The dose-dependent anti-inflammatory properties that oleocanthal exhibits in vitro are mimicked by its dose-dependent irritation in the oral cavity (Beauchamp et al., 2005), making oral irritation (measured by the taste bioassay) a de facto marker of biological activity (Cicerale, Breslin, Beauchamp, & Keast, 2009; Fischer, Griffin, Archer, Zinsmeister, & Jastram, 1965; Joshi, Hankey, & Patwardhan, 2007). Moreover, the transient receptor potential cation channel A1 (TRPA1) has been identified as being the sensory receptor linked to oleocanthal, and the anatomical location of the TRPA1 channel specific to this compound has been found in the oropharyngeal region of the oral cavity (Peyrot des Gachons et al., 2011). Immediately following oil extraction from the olive fruit, there is potential for the phenolic quality of the oil to decline, via oxidation catalysed by oxygen (O2) and light (Brenes, Garcia, Garcia, & Garrido, 2001; Kalua, Bedgood, Bishop, & Prenzler, 2006; Morello, 2004). To preserve the quality of EVOO and consequently the concentration of beneficial phenolic compounds in such oils, we need to understand the extent of degradation caused by O2 and light (Kalua et al., 2006). Oxidation that occurs in edible oils relates to the loss of minor components and formation of new compounds, causing nutritional loss as well as the development of rancid and off-flavours. Autoxidation and photosensitised (light) oxidation of lipids can occur through the interaction of fatty acids with triplet and singlet oxygen. It can also occur as a result of the degradation of hydroperoxides stimulated by light or the presence of trace transition metals. In the process of autooxidation and photosensitised oxidation, olive oil phenolic compounds are generally degraded, causing the loss of health benefiting compounds (Lerma-Garcia, Herrero-Martinez, Simo-Alfonso, Lercker, & Cerretani, 2009). To date, four studies have examined the influence of light over time on changes in concentration of phenolic compounds in EVOO. These investigations demonstrated a decrease in the concentration of phenolics upon light exposure over an extended period of storage (Gutierrez & Fernandez, 2002; Luna, Morales, & Aparicio, 2006; Okogeri & Tasioula-Margari, 2002; Rastrelli, Passi, Ippolito, Vacca, & De Simone, 2002). One study in particular by Okogeri and TasioulaMargari (2002) showed that there was a 57–63% reduction in total phenolic content upon six months of EVOO storage under diffused light. For the EVOO stored in darkness there was a smaller reduction in total phenolic content (39–45%). In general though, oxidation in EVOO is an inevitable process of ageing and therefore cannot be fully prevented (even without exposure to such a factor as light) (Kristott, 2000). Several studies have examined changes in phenolic concentrations upon storage without exposure to light (Baiano et al., 2009; Brenes et al., 2001; Cinquanta, Esti, & La Notte, 1997; Gomez-Alonso, Manebo-Campos, Desamparados Salvador, & Fregapane, 2007; Lavelli, Fregapane, & Salvador, 2006; Lerma-Garcia, Herrero-Martinez, et al., 2009; LermaGarcia, Simo-Alfonso, et al., 2009; Morello, 2004; Rastrelli et al., 2002). It is important to note that one study (Lerma-Garcia, Simo-Alfonso, et al., 2009), stored EVOO in uncapped bottles (therefore allowing O2 to filter through freely) and the remaining studies did not indicate if the oils were kept under O2 free conditions (Baiano et al., 2009; Brenes et al., 2001; Cinquanta et al., 1997; Gomez-Alonso et al., 2007; Lavelli et al., 2006; Morello, 2004). Overall, these studies demonstrated significant decreases in phenolic concentration upon extended storage even without light exposure. EVOO generally has a relatively long shelf-life of approximately 12–18 months (Morello, 2004). This time frame for shelf-life is appropriate as olive fruit used for olive oil production is harvested annually (Boskou, 2006). For oleocanthal to indeed provide health benefits, it must be present in significant quantities throughout the shelf-life of EVOO. Therefore, investigation into the stability of

oleocanthal upon extended storage and exposure to both light and O2 (via atmospheric air) as would occur in a common domestic situation is warranted. Hence, the aim of the current study was to investigate the extent of degradation of oleocanthal in EVOO when exposed to natural light and O2 over time. 2. Materials and methods 2.1. Oil samples One EVOO (Red Island Australia, Australia) containing 90 mg/kg oleocanthal was stored for 10 months with and without exposure to light (approximately 1824 h, this was based on an average of 6 h of sunlight per day in Melbourne as stated by the Australian Government Bureau of Meteorology) and O2 via natural air exposure. The experimental design was full factorial and there were four differing treatments: oil not exposed to light and no (or minimal) exposure to O2 (NLNO2), oil exposed to O2 only (NLYO2), oil exposed to light only (YLNO2) and oil exposed to both light and O2 (YLYO2). Oil without exposure to light and minimal exposure to O2 was stored in darkness in a nitrogen (N2) purged capped bottle wrapped in aluminium foil. Oil exposed to O2 only, was stored in darkness in an uncapped bottle, wrapped in aluminium foil. Oil exposed to light only was stored in a N2 purged capped clear bottle, in a naturally lit room near a window. Oil exposed to both light and O2 was stored in a clear glass bottle in a naturally lit room near a window, uncapped. All oils were stored at room temperature (20 ± 3 °C). For each treatment, 4.5 L of oil was stored in a 5 L schott bottle. Bottles were stirred prior to samples being removed (140 ml in total, 70 ml for HPLC analysis and 70 ml for biological activity analysis at any given time point) with a pipette. Analysis of oils occurred once per month for the first six months of storage and then a final analysis was conducted at the 10 month time point. Oleocanthal concentration was determined by calculating the ratio of oleocanthal peak area to 3, 5-dimethoxyphenol (internal standard [ISTD]) peak area and multiplied by the amount of ISTD added to the sample (Cicerale, Conlan, Barnett, Sinclair, & Keast, 2009). For the additional phenolics quantified (oleuropein, deacetoxy oleuropein aglycone, oleuropein aglycone, luteolin and hydroxytyrosol), concentration was calculated in the same way as for oleocanthal. A negative control—corn oil (devoid in oleocanthal) (Nature First, Australia) was also included in the study. 2.2. High performance liquid chromatography analysis The method used for preparation and analysis of oleocanthal in olive oil was modified from the method developed by Beauchamp et al. (2005), Impellizzeri and Lin (2006) and more recently Cicerale et al. (2009). Samples were prepared and analysed in triplicate. A stock solution (5000 mg/kg) of the ISTD, 3, 5-dimethoxyphenol (Aldrich, U.S.A) was prepared in methanol (Rowe Scientific, Australia). This solution was spiked in the oil samples. To quantify oleocanthal using internal calibration, the ISTD was added to the oil (10 g) at a concentration of 50 mg/kg. Oleocanthal concentration was expressed as 3, 5-dimethoxyphenol (ISTD) equivalents. Oleocanthal was extracted from the oil matrix by liquid–liquid partitioning according to the following procedure. Hexane (5 ml) (Rowe Scientific, Australia) was added to a centrifuge tube containing 10 g of oil. The tube was vortexed for 1 min to mix oil and hexane thoroughly. Acetonitrile (30 ml, extraction solvent) (Rowe Scientific, Australia) was added and then vortexed for 1 min. The tube was centrifuged at 4000 rpm for 5 min to separate the solvent from the oil phase. The solvent extract was collected and placed in an evaporator flask and the solvent was removed with use of a rotary evaporator (Buchi, Switzerland) at 45 °C and 200 Mbar. Methanol–water (1 ml) (HPLC grade) (1/1, v/v) was pipetted into the flask containing the dried down extract to dissolve the residue of this extract. The extract was then

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transferred to a centrifuge tube. Hexane (1 ml) was added to the solution to aid in the removal of any residual oil. The tube was vortexed for 1 min and then centrifuged (4000 rpm for 2 min) for phase separation. The methanol–water phase was collected and filtered (with the use of a 0.45 μm filter) (Phenex, U.S.A) for HPLC analysis. Oleocanthal was separated using a 1200 series HPLC system with solvent degasser, quaternary pump, auto sampler and diode array detector set to 278 nm (Agilent Technologies, Blackburn, Australia). An Apollo RP-C18 column (250 mm × 4.6 mm ID, 5 μm; Grace Davison, Baulkham Hills, Australia) was used for all separations at a constant temperature of 25 °C using the gradients previously described (Cicerale, Conlan, Barnett, et al., 2009), at a flow rate of 1 ml/min with an injection volume of 20 μl. Oleocanthal was identified using a 6210 MSDTOF mass spectrometer (Aglient Technologies, Blackburn, Australia) under the following conditions: drying gas, nitrogen (7 ml− 1, 350 °C); nebuliser gas, nitrogen (15 psi); capillary voltage 4.0 kV; vaporiser temperature 350 °C; and cone voltage 60 V. 2.3. Biological activity analysis The method used for the oleocanthal biological activity assay was adapted from the taste bioassay protocol developed by Beauchamp et al. (2005) and more recently Cicerale et al. (2009). Oleocanthal elicits irritation in the oral pharynx in a dose-dependent manner, mirroring COX inhibitory activity meaning biological activity can be determined via a taste bioassay (Beauchamp et al., 2005). Subjects (n = 11, 34.6 ± 10.7 years, 8 women and 3 men) between the ages of 23 and 52 years were university staff and students in Melbourne, Australia. Originally 14 people were recruited however 3 subjects were unable to complete the study. All subjects agreed to participate in the study and provided informed consent on an Institutional Review Board form approved by the Deakin University Ethics Committee (EC253—2006). The participants, were requested to refrain from consuming food and drink (except room temperature water) and using oral irritants (such as toothpaste, mouthwash and gum) 2 h prior to testing. Participants were trained in the use of the general Labelled Magnitude Scale (gLMS) following the published procedures by Bartoshuk et al. (Bartoshuk et al., 2004; Green et al., 1996; Green, Shaffer, & Gilmore, 1993; Keast & Roper, 2007). Participants attended a training session to evaluate oleocanthal irritation intensity with the use of an EVOO containing oleocanthal. All testing took place in a specialised testing facility comprising seven individual computerised booths, containing red lights to mask potential colour differences in the oils. Each subject was isolated from other subjects by vertical dividers to eliminate interaction between subjects (Keast & Roper, 2007). Test sessions were scheduled so that there was a minimum of at least 2 h between each test. Each sample was evaluated in duplicate. Stimulus delivery was the same as used by Cicerale et al. (2009). Briefly, an aliquot of 5 ml of each oil (n = 20) was presented in 30 ml polyethylene medicine cups (McFarlane Medical, Surrey Hills, Australia) in a fully randomised order. Subjects rinsed their mouths with filtered (fi) water (8 μm particulate filter with an activated charcoal filter, Dura®) at least three times over a 2 min period before commencement of testing. Subjects wore nose clips to eliminate olfactory cues. Each subject swallowed the oil in two aliquots and after 20 s had elapsed, subjects were asked to rate (with use of the gLMS) the oils for intensity of peak throat irritation. 2.4. Data analysis Data was analysed using SPSS for Windows, version 17.0 (SPSS 17.0, Chicago, Illinois). Statistical comparison of means was investigated with a two-way ANOVA and Tukey post-hoc analysis. P values of b0.05 were

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considered to be significant. A Pearson product–moment correlation was also conducted to investigate the relationship between the HPLC quantified concentration and biological activity of oleocanthal in the EVOO samples. 3. Results 3.1. Chemical stability of oleocanthal upon storage over time and under different storage conditions A two-way ANOVA revealed there was a significant effect of time [F (5, 50) = 28.66, p b 0.05] on oleocanthal concentration for all the oils when combined. Post-hoc analysis revealed that there were significant differences (p b 0.05) between: month 1 and months 6 and 10; month 2 and all other months (except month 3); month 3 and months 6 and 10; month 4 and months 2, 6 and 10; month 5 and months 2, 6 and 10; month 6 and all other time points (except month 10); and month 10 and all other time points (except month 6). A one-way ANOVA on each treatment demonstrated that there were significant (p b 0.05) differences in oleocanthal concentration over time. Refer to Table 1 for mean values ± SD. A two-way ANOVA also revealed that there was a significant difference amongst the different storage treatments [F (3, 48) = 2.72, p = 0.05]. Further oneway ANOVA analysis revealed that across treatments, YLYO2 was significantly different to NLNO2 at month 10 (p b 0.05). The overall change in oleocanthal concentration between the different oils (over the 10-month storage period) was as follows: 15% (90±13 mg/ kg to 76±9 mg/kg), 28% (90±13 mg/kg to 64±4 mg/kg), 25% (90± 13 mg/kg to 67±5 mg/kg) and 37% (90±13 mg/kg to 56±9 mg/kg) respectively for the treatments NLNO2, NLYO2, YLNO2 and YLYO2. As for the additional phenolics quantified, a similar pattern of degradation was noted (refer to Fig. 1). Firstly oleuropein degradation was as follows: 27%, 63%, 38% and 82% for NLNO2, NLYO2, YLNO2 and YLYO2 respectively. Secondly deacetoxy oleuropein aglycone degradation was: 27%, 70%, 41% and 77% for NLNO2, NLYO2, YLNO2 and YLYO2 respectively. Thirdly, oleuropein aglycon degradation was: 1%, 46%, 71%, 73% respectively. Fourthly, luteolin degradation was: 16%, 39%, 35%, 68% respectively. Contrary to the phenolics discussed thus far, an increase in hydroxytyrosol (40–69%) was found after 10 months storage. 3.2. Stability of oleocanthal biological activity upon storage over time and under different conditions A two-way ANOVA revealed there was a significant effect of time [F (5, 453) = 12.06, p b 0.05] and treatment [F (3, 453) = 22.15, p b 0.05] on oleocanthal biological activity. Further post-hoc analysis revealed that these significant differences (p b 0.05) were between month 10 and all other time points (except month 1). It was also found that oleocanthal biological activity for the oil, YLYO2 significantly differed (p b 0.05) from that of treatments NLNO2, NLYO2 and YLNO2. Further one-way ANOVA analysis on each treatment demonstrated significant (p b 0.05) differences in oleocanthal concentration over time. Refer to Table 2 for mean values ± SD. 4. Discussion Oleocanthal concentration decreased somewhat (15–37%) over a 10 month storage period, depending on the storage conditions. This data supports previous research in that, phenolic compounds of secoiridoid structure generally breakdown and decrease in concentration over the course of storage (Baiano et al., 2009; Brenes et al., 2001; Lerma-Garcia, Herrero-Martinez, et al., 2009; Lerma-Garcia, Simo-Alfonso, et al., 2009; Morello, 2004; Stefanoudaki, Williams, & Harwood, 2010). The overall change in oleocanthal concentration between the different treatments was as follows: 15%, 28%, 25% and 37% for NLNO2, NLYO2, YLNO2 and YLYO2, respectively. In considering the influence of

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Table 1 Oleocanthal concentration (ISTD equivalents) quantified by HPLC (mean ± SD) for the control and oil treatments: NLNO2, NLYO2, YLNO2 and YLYO2 during storage. August (control, month 1) Oil treatment September (month 2) October (month 3) November (month 4) December (month 5) January (month 6) May (month 10) 90 ± 13abc q 90 ± 13defg q 90 ± 13ijk q 90 ± 13no q

NLNO2 NLYO2 YLNO2 YLYO2

107 ± 2ab r 104 ± 12deg r 84 ± 3ijkl r 89 ± 15no r

85 ± 7ac s 89 ± 3defg s 91 ± 1ijk s 86 ± 4no s

85 ± 5ac t 78 ± 3dfgh t 79 ± 2ijklm t 88 ± 10no t

83 ± 9ac u 83 ± 3defgh u 84 ± 4ijkl u 86 ± 6no u

74 ± 1ac v 70 ± 9dfgh v 71 ± 6jklm v 71 ± 9nop v

76 ± 9ac wx 64 ± 4fgh wx 67 ± 5klm wx 56 ± 9op x

Mean values with unlike superscripts were significantly different (p b 0.05). Superscripts (a–p) depict significant differences for each treatment at the differing storage times. Superscripts (q–x) depict significant differences between each treatment.

light and O2 exposure, both factors appear to be important in reducing oleocanthal concentration and biological activity during storage. For instance, oleocanthal concentration decreased 28% in the oil exposed to O2 but not light (NLYO2) over the course of 10 months. For the oil not exposed to O2 but exposed to light (YLNO2), oleocanthal concentration had decreased at a similar level of 25%. Data on the additional phenolics quantified (oleuropein, deacetoxy oleuropein aglycone, oleuropein aglycone, luteolin and hydroxytyrosol) also supports earlier research. Research has shown that phenolic compounds of secoiridoid structure generally breakdown and are lost over the course of storage (Baiano et al., 2009; Brenes et al., 2001; Lerma-Garcia, Herrero-Martinez, et al., 2009; Lerma-Garcia, SimoAlfonso, et al., 2009; Morello, 2004; Stefanoudaki et al., 2010). With regards to hydroxytyrosol, a rise in this compound's concentration over a period of storage has been previously noted (Baiano et al., 2009; Brenes et al., 2001; Morello, 2004; Stefanoudaki et al., 2010). Baiano et al. (2009) concluded that the decrease in secoiridoid phenolics and simultaneous increase in phenolic alcohols (such as hydroxytyrosol) over storage was a consequence of ligstroside and oleuropein compounds degrading, which in turn act as a substrate for the formation of such phenolic alcohols.

The detrimental effect of O2 on oleocanthal concentration may be a result of not only O2 causing oxidation of phenolics but possibly the activation of polyphenoloxidase (PPO) and peroxidase (POD) enzymes also. PPO and POD become activated in the presence of O2 and possess the ability to oxidise phenolic compounds, resulting in a reduction in their concentration (Migliorini, Mugelli, Cherubini, Viti, & Zanoni, 2006). Thus, increased oil exposure to O2 may have stimulated increased activity of these enzymes. Vierhuis et al. (2001) demonstrated that EVOO produced using N2 purging (thus eliminating O2) during the process of malaxation, inhibited PPO and POD activities and resulted in an increase in phenolic concentration. Hence, minimisation of O2 exposure during production and storage of EVOOs appears an imperative measure to maximise phenolic retention, including oleocanthal. It is also evident that light exposure is detrimental to oleocanthal. This damaging effect may occur via photosensitised oxidation of lipids through interaction of fatty acids with triplet and singlet oxygen or degradation of hydroperoxides (Lerma-Garcia, Herrero-Martinez, et al., 2009; Lerma-Garcia, Simo-Alfonso, et al., 2009). As with O2, light exposure to EVOO should be avoided. If an olive oil consumer ingests around 50 g of EVOO a day that contains approximately 100 mg/kg of oleocanthal, the person would

Fig. 1. Phenolic concentrations including oleocanthal, for the treatments: (a) NLNO2, (b) NLYO2, (c) YLNO2 and (d) YLYO2 at T1 (beginning of storage) and T2 (end of 10 months storage).

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Table 2 Oleocanthal biological activity (gLMS units) quantified by the taste bioassay assay (mean ± SE) for the oil treatments: NLNO2, NLYO2, YLNO2 and YLYO2 during storage (n = 11). August (control, month 1) Oil treatment September (month 2) October (month 3) November (month 4) December (month 5) January (month 6) May (month 10) 20 ± 0.5ab p 20 ± 0.5cdeg p 20 ± 0.5hi p 20 ± 0.5lmno p

NLNO2 NLYO2 YLNO2 YLYO2

22 ± 0.4b q 24 ± 0.7cdefg q 22 ± 0.3ij q 23 ± 0.2lmo q

26 ± 0.2ab rs 23 ± 0.3cdefg rst 24 ± 0.05ij rst 22 ± 0.3lmno st

23 ± 0.9ab u 25 ± 0.01cdef u 24 ± 0.06ij u 23 ± 0.2lmo u

22 ± 0.7ab v 24 ± 0.4cdef v 24 ± 0.2ij v 19 ± 0.2ln v

25 ± 0.3ab w 26 ± 0.4def w 23 ± 0.2hij w 23 ± 0.2lmo w

23 ± 0.03ab xz 19 ± 0.3cdg xz 19 ± 0.4 z 6 ± 0.2

Mean values with unlike superscripts were significantly different (p b 0.05). Superscripts (a–o) depict significant differences for each treatment at the differing storage times. Superscripts (p–z) depict significant differences between each treatment at one given time.

consume approximately 5 mg/kg a day of oleocanthal. Using the current data to calculate the effect on oleocanthal intake, if 50 g of EVOO exposed to both light and O2 (YLYO2) containing 56 mg/kg of oleocanthal was ingested, this would correspond to an intake of 2.8 mg/d. If 50 g of EVOO not exposed to light and minimal O2 (NLNO2) containing 76 mg/kg of oleocanthal was ingested, this would correspond to a slightly higher intake of 3.8 mg/d. Overall, oleocanthal appears to be present in potentially pharmacologically significant quantities after an extended storage period and upon being exposed to known oxidative agents. Historically, olive oil is an annual product therefore the health benefiting compounds should not completely degrade over a 12 month period. If oleocanthal was to degrade extensively with storage and upon exposure to light and O2, then it would more than likely not act as a health benefiting compound, as it would not be present in the oil. As expected, oleocanthal biological activity data was generally in agreement with the HPLC data obtained in that, biological activity remained fairly stable over storage. A correlation analysis was conducted and demonstrated that a strong positive correlation (r = 0.8, p b 0.05) existed between the HPLC analysis of concentration and the biological activity of oleocanthal. However, it is important to note that after 10 months storage, biological activity of the oil, YLYO2 reduced at a greater increment (70%) than that of actual concentration (37%). This supports earlier research which showed a greater decrease in oleocanthal biological activity (31%) compared to absolute oleocanthal concentration (16%) after an EVOO was heated for an extended period of time (Cicerale, Conlan, Barnett, et al., 2009). A possible explanation for this finding is that during the degradation process new compounds (an oleocanthal antagonist) may form, decreasing or masking the biological activity of oleocanthal. Another plausible explanation is that the human sensory system may be sensitive to a modest decrease in oleocanthal concentration. The oral dose–response curve of compounds (Ψ = k Φn) is rarely a 1:1 ratio or liner relationship (Meilgaard, Civille, & Carr, 1999). When n is greater than one, the sensation grows faster than the stimulus and when n is smaller than one the sensation grows more slowly. For example, the exponent of oral activation by quinine–HCl and NaCl is 0.65 and 1.4 respectively. This means that an increase of 1 mM quinine–HCl would result in a 0.65 increase in oral perception of this compound, whereas an increase of 1 mM of NaCl would result in a 1.4 increase in oral perception of NaCl (Meilgaard et al., 1999). Also, while it is highly unlikely given previous published data (Beauchamp et al., 2005; Peyrot des Gachons et al., 2011), we should not discount that other phenolic compounds may have a synergistic effect on oropharyngeal irritation and degradation of those compounds may result in decreased irritation. In conclusion, oleocanthal concentration degraded somewhat upon extended storage (up to 37%) however, the data still demonstrated a potentially significant health promoting pharmacologic capacity after 10 months. This data provides further evidence that oleocanthal is a health benefiting agent, potentially contributing to the beneficial effects of a traditional Mediterranean diet, which includes the consumption of large volumes of annually produced EVOO. If oleocanthal was to degrade extensively with storage and upon exposure to light and O2, then it would more than likely not act as a health benefiting compound, as it would not be present in the oil.

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