Physical Stability, Autoxidation, and Photosensitized Oxidation of Ï-3

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Physical stability, autoxidation and photosensitized oxidation of #-3 oils in nanoemulsion prepared with natural and synthetic surfactants Sibel Uluata, D. Julien McClements, and Eric A. Decker J. Agric. Food Chem., Just Accepted Manuscript • DOI: 10.1021/acs.jafc.5b03572 • Publication Date (Web): 09 Oct 2015 Downloaded from http://pubs.acs.org on October 10, 2015

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Journal of Agricultural and Food Chemistry

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Physical stability, autoxidation and photosensitized oxidation

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of ω-3 oils in nanoemulsion prepared with natural and

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synthetic surfactants Sibel Uluata*,a,b, D. Julian McClementsa,c, Eric A. Deckera,c

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5 6

a

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Laboratory 100 Holdsworth Way Amherst, MA 01003 USA

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b

Department of Food Technology, Inonu University, 44280 Malatya, Turkey

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c

Department of Biochemistry, Faculty of Science, King Abdulaziz University, P. O. Box

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Department of Food Science University of Massachusetts Amherst 228 Chenoweth

80203 Jeddah 21589 Saudi Arabia

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ACS Paragon Plus Environment


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Abstract

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The food industry is interested in the utilization of nanoemulsions stabilized by natural

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emulsifiers little research has been conducted to determine the oxidative stability of such

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emulsions. In this study, two natural (lecithin and quillaja saponin) and two synthetic

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(Tween 80 and sodium dodecyl sulfate) surfactants were used to fabricate omega-3

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nanoemulsion using high pressure homogenization (microfluidization). Initially, all the

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nanoemulsions contained small (d from 45 to 89 nm) and anionic (ζ–potential from -8

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and –65 mV) lipid droplets (pH 7). The effect of pH, ionic strength and temperature on

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the physical stability of the nanoemulsions system was examined. Nanoemulsion

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stabilized with Tween 80, quillaja saponin or sodium dodecyl sulfate (SDS) exhibited no

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major changes in particle size or visible creaming in the pH range of 3 to 8. All

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nanoemulsions were relatively stable to salt addition (0 to 500 mM NaCl, pH 7.0).

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Nanoemulsion stabilized with SDS and quillaja saponin were stable to heating (30 to 90

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o

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absence of the singlet oxygen photosensitizers, riboflavin and rose bengal. Riboflavin

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and rose bengal accelerated lipid oxidation when compare to samples without

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photosensitizers. Lipid hydroperoxide formation followed the order Tween 80 > SDS >

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lecithin > quillaja saponin and, propanol formation followed the order lecithin > Tween

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80 > SDS > quillaja saponin at 37 oC for autoxidation. The same order of oxidative

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stability was observed in the presence of photosensitize oxidation promoted by riboflavin.

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Quillaja saponin consistently produced the most oxidatively stable emulsions, which

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could be due to its high free radical scavenging capacity.

C). The impact of surfactant type on lipid oxidation was determined in the presence and


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Key words: nanoemulsions, omega-3 oil, photosensitized oxidation, surfactant, Quillaja

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saponin, Riboflavin, Rose bengal, lipid oxidation

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INTRODUCTION

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(DHA) and eicosapentaenoic acid (EPA).(1) These fatty acids are important for proper

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brain development and the reduction of cardiovascular diseases and inflammation.(2-4)

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Omega-3 fatty acids are currently under consumed by many individuals so there is

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interest in omega-3 fortified foods.(5) The major sources of omega-3 fatty acids (such as

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fish, algae, and flaxseed oils) are highly hydrophobic molecules with low water solubility

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making them difficult to incorporate into aqueous based foods.(4) They are also prone to

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chemical degradation due to lipid oxidation, which reduces product quality due to the

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production of undesirable off-flavors.(6) Consequently, there is a need to develop

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effective delivery systems that both incorporate omega-3 oils into water based food and

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protect them from oxidation so they can be used to fortify commercial foods and

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beverages.

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Oil-in-water nanoemulsions are delivery systems that contain lipid droplets with a mean

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particle diameter below 200 nm.(7) When the lipid droplet dimensions become

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appreciably smaller than the wavelength of light (d < λ/20), the light waves are only

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scattered weakly thereby giving nanoemulsions a transparent or slightly turbid

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appearance.(8) Optical clarity is an important attribute for delivery systems that need to be

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incorporated into transparent food and beverage products, such as some fortified waters,

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soft drinks, dressings, and sauces.(9, 10)

Fish oil is an important source of the omega-3 fatty acids, docosahexaenoic acid

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The small size of the droplets in nanoemulsions means that they typically have

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higher physical stability than conventional emulsions, and are more effective at


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enhancing the bioavailability of encapsulated lipids.(11) Nevertheless, there may also be

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some drawbacks associated with using nanoemulsions as delivery systems for omega-3

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fatty acids. First, the small droplet size means that there is a high droplet surface area,

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which could promote lipid oxidation since this reaction usually occurs at the oil-water

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interface.(12) Second, the increased transmission of light waves through nanoemulsions

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could increase their susceptibility to light-induced oxidation. Consequently, there is a

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need for research to understand the influence of nanoparticle characteristics on lipid

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oxidation in nanoemulsions before they can successfully be utilized as delivery systems.

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Light exposure is one of the important factors promoting lipid oxidation in foods.(13)

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Unsaturated fatty acids do not strongly absorb the relatively low energy waves of visible

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light, but they do absorb the relatively high-energy waves of ultraviolet light.(13,

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Consequently, lipids normally tend to be fairly stable to oxidation in the presence of

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visible light, but oxidize rapidly in the presence of UV light. However, photosensitizers

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such as chlorophylls, riboflavin and rose bengal can be excited by visible light and

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produce reactive oxygen species that are capable of accelerating lipid oxidation.(15-17)

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Photosensitizer-induced oxidation can take place by two different pathways (Type I and

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II) depending on the photosensitizer type and environmental conditions such as oxygen

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concentration, solvent type and aggregation of molecules.(14) In both pathways, light

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waves convert the photosensitizer from a ground state (singlet) to an excited state

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(triplet). In type I pathway, the excited triplet state of the photosensitizers can directly

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abstract electrons or hydrogen atom from substrates such as polyunsaturated fatty acids to

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generate free radicals. In type II pathway, the excited triplet photosensitizers transfer their

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energy to triplet oxygen to generate singlet oxygen.(15, 17-20) Singlet oxygen can then


14)


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directly attach to the double bonds of unsaturated fatty acids to produce lipid

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hydroperoxides. Previous studies showed that riboflavin photosensitization produces

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both singlet oxygen and superoxide anion in oil-in water emulsions, thereby promoting

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lipid oxidation.(19) These researchers proposed that singlet oxygen produce by the type II

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pathway can react with unsaturated fatty acid to generate hydroperoxides. In addition, the

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superoxide anion produced by photosensitized riboflavin could reduce ferric (Fe3+) to

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more reactive ferrous (Fe2+) ions which in turn could accelerate the decomposition of

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hydroperoxides produced by singlet oxygen.(19) Overall, the potential of lipids to be more

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susceptible to oxidation in nanoemulsions needs to be understood if the food and

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pharmaceutical industries are to use these delivery systems in their products. (9, 21, 22)

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Emulsifier type can have a major impact on lipid oxidation especially when prooxidants

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exist in the aqueous phase e.g., transition metals(23-26) and thus the emulsion droplet

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interface can influence prooxidant-lipid interactions. For example, negatively charged

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emulsions droplets are more susceptible to metal promoted lipid oxidation than positively

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charged emulsion droplets since metals are attracted to the anionic emulsion droplet

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interface where they can readily interact with lipids in the emulsion droplet core.(27)

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The food and beverage industry is trying to replace synthetic surfactants with more

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natural “label-friendly” ingredients.

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were to determine how synthetic (Tween 80 and SDS) and natural (lecithin and quillaja

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saponin) surfactants impacted the physical stability of nanoemulsions, autoxidation and

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photosensitized lipid oxidation in oil-in-water emulsions. Since some foods are

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susceptible to light promoted oxidation, oxidation studies were done in the absence and

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presence of the photosensitizer, riboflavin. Commercially available lecithin and quillaja

(28)

Therefore, the overall objectives of this study


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saponin (Q-naturale) was used to better understand the stability of the nanoemulsions

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prepared with commercial emulsifiers. Fish oil ethyl esters were used as the lipid source

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as they represent a highly concentrated form of omega-3 fatty acids that could provide

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useful for transparent food applications where minimizing total lipid concentration

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improves clarity.

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MATERIALS AND METHODS

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Materials

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Fish oil ethyl ester (MEG-3 3327 EE) was kindly provided by DSM Nutritional

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Products Ltd. (Basel, Switzerland) and contained 55 % omega-3 fatty acids as determined

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by the manufacturer. Lecithin (Sunlipon 65) were provided by Perimondo (Newyork,

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USA). Sunlipon 65 contained 64% phosphotidylcholine. Q-Naturale 200 was provided

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from Ingredion Inc. (Westchester, Il) and contained 14% quillaja saponin in water as

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indicated by the manufacturer. Polyoxyethylene (20) sorbitan monooleate (Tween 80),

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sodium dodecyl sulfate (SDS >98%), riboflavin, rose bengal, AAPH (2,2′-azobis(2-

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amidinopropane) dihydro-chloride) and Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-

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2-carboxylic acid) were purchased from Sigma-Aldrich (St. Louis, MO, USA). All other

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reagents were of analytical or chromatographic grade.

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Emulsion preparation

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Oil-in-water coarse emulsions were prepared by homogenizing 1 wt % fish oil ethyl

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esters with 99 wt % aqueous phase. The aqueous phase consisted of surfactant (1.5 wt %

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Tween 80, SDS, lecithin or quillaja saponin) and 10 mM sodium phosphate buffer

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solution (pH 7). A coarse emulsion was prepared by blending the lipid and aqueous



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phase using a high-shear mixer (M133/1281-0, Biospec Products, Inc., Bartlesville, OK)

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for 2 min. The coarse emulsions were then homogenized using a microfluidizer (M-110P,

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Microfluidics, Newton MA, USA) using 5 passes at 20,000 psi to produce nanoemulsions.

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Particle size and ζ-potential measurements

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The particle size distributions and droplet charge (ζ-potential) of the nanoemulsions

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were measured using a particle electrophoresis instrument (Zetasizer a Nano ZS series

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Malvern Instruments, Worchester, UK). Samples were diluted 1:100 times using a 10

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mM phosphate buffer solution (pH 7) before analysis to prevent multiple scattering. The

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results of particle size are expressed as the Z-average particle diameter.

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Emulsion stability testing

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The stability of the nanoemulsion was determined by measuring changes in particle size

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and making visual observation of creaming under a variety of environmental conditions

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during storage at room temperature for 24 h. The effect of pH was determined over the

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range of 2 to 8 by adjusting pH of the emulsions using 1 M HCl and NaOH. The effect

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of salt addition was determined by adding different amounts of 1 M NaCl solution to the

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nanoemulsions to reach final concentrations of 0 to 500 mM (pH 7.0). The influence of

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heating was determined by placing the nanoemulsions into a water bath at 30 to 90 oC for

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30 min followed by immediate cooling at room temperature.

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Absorbance measurements

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The absorbance spectra of individual surfactants (Tween 80, SDS, lecithin, quillaja

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saponin), riboflavin and rose bengal was measured using with a UV-visible


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spectrophotometer (UV-2101 PC, Shimadzu Co., Tokyo, Japan) at a wavelength from

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320 to 700 nm.

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ORAC Assay

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The free radical scavenging activity of surfactants was measured using the oxygen

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radical absorbance capacity assay (ORAC).(29)

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automated plate reader (Synergy HT Multi-Mode Microplate Reader; BioTek

163

Instruments, Winooski, VT) with 96-well plates. The excitation wavelength was 485 nm

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and the emission wavelength was 528 nm. All reagents were prepared in 75 mM

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phosphate buffer solution (pH 7.4). Stock solutions of fluorescein (93.5 nM), AAPH

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(221mM) and Trolox (standard, 0-100 µM) were prepared daily. To each well, 50 µl of

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fluorescein, 50 µl of buffer and 50 µl of sample (0-100 µM) or Trolox was added. The

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plate were preheated for 5 min at 37°C prior to the addition of 50 µl AAPH. Immediately

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after the addition of AAPH, fluorescence was read every min for 2 h. ORAC values were

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expressed as µmole of Trolox equivalents (TE) per µmole of surfactant using a Trolox

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standard curve prepared with 0 and 100 µM Trolox.

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Lipid oxidation studies

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Each nanoemulsion sample (1 mL) was put in a 10 mL vial and sealed airtight. Sample

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vials were incubated at 37 under 1440-lux light intensity (measured by a Sunche light

175

meter model HS1010) or in the dark. For photosensitized studies, riboflavin and rose

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bengal were added to the nanoemulsions at a final concentration of 5 µM.

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Lipid oxidation was monitored with hydroperoxides and propanal. Lipid hydroperoxide

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were determined according to Uluata et al.(30) Emulsion samples (0.3 mL) were mixed

Assays were performed with an



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with 1.5 ml of isooctane/2-propanol solution (3:1 v/v) and vortexed (10 s, 3 times). The

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mixed solution was centrifuged at 3400g for 10 min (Centrific TMCentrifuge, Thermo

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Fisher Scientific Inc., Fairlawn, NJ, USA). The upper organic layer (200 µL) was mixed

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with 2.8 ml of methanol/butanol solution (2:1, v/v), followed by the addition of 15 µL of

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3.94 M ammonium thiocyanate and 15 µL of Fe2+solution. The Fe2+ solution was

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prepared freshly from the supernatant of a mixture of equal amounts of 0.132 M BaCl2 in

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0.4 M HCl and 0.144 M FeSO4. The solution was vortexed, held 20 min at room

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temperature and the absorbance was measured at 510 nm in a UV–Vis spectrophotometer

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(Genesys 20, Thermo Fisher Scientific Inc., Waltham, MA, USA). Hydroperoxide

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concentrations were determined using a standard curve prepared from cumene

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hydroperoxide. Each measurement was performed in triplicate and results were expressed

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as mean values ± standard deviation.

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Headspace propanal was measured according to the method described with minor

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modification by Panya et al(31) using a gas chromatograph (Model GC-2014, Shimadzu

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Co., Tokyo, Japan) equipped with a solid phase micro extraction (SPME) auto injector

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(Model AOC-5000, Shimadzu Co., Tokyo, Japan). Emulsions (1 mL) in 10 ml glass vials

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capped with aluminum caps with polytetrafluoroethylene (PTFE)/Silicone septa were

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heated at 45 0C for 10 min in a heating block before measurement. A 50/3 µm

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divinylbenzene/carboxen/polydimethylsiloxane

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(SPME) fiber (Supelco Co., Bellefonte, PA, USA) was then inserted into the vial

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headspace for 2 min to absorb volatiles. The fiber was transferred to the GC injector port

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(250 0C) for 3 min. The injection port was operated at a split ratio of 1:7. Volatiles were

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separated on a fused-silica capillary Equity-1 Supelco column (30 mm x 0.32 mm i.d.x

(DVB/Carboxen/PDMS)


stable

flex

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µm) coated with 100% poly-dimethylsiloxane (PDMS) at 70 0C for 10 min. A flame

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ionization detector was used at a temperature of 250 0C. Propanal concentrations were

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determined from peak areas using a standard curve made from authentic propanal.

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2.6. Statistical Analysis

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All experiments were analyzed in triplicate and repeated twice and results expressed

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as mean values ± standard deviation. All data result were analyzed by analysis of

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variance (ANOVA) using SPSS 20 (SPSS Inc.,Chicago, IL). The differences between

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mean values were compared using Duncan's multiple-range test with level of significance

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of p ≤ 0.05.

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RESULTS AND DISCUSSION

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Physical stability of the nanoemulsions

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Initially, the particle size and charge of the lipid droplets in the nanoemulsions

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prepared using different natural and synthetic surfactants were measured. The mean

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particle diameter of all the samples was lower than 100 nm, confirming that these

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delivery systems were nanoemulsions. The relatively small droplet size can be attributed

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to the relatively large surfactant-to-oil ratio (SOR = 1.5) used to produce the

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nanoemulsions. Nevertheless, there were appreciable differences in the mean droplet

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diameters produced in the different nanoemulsions: Tween 80 =45 nm; SDS = 67 nm;

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lecithin = 82 nm and quillaja saponin = 89 nm (Table 1). All the nanoemulsions were

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slightly turbid in appearance (Figure 1).

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At pH 7.0, all the lipid droplets in the nanoemulsions were negatively charged with

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the magnitude of the negative charge decreasing in the following order: Tween 80
lecithin>

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quillaja saponin. Exposure to light only increased the formation of lipid hydroperoxides

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in the Tween 80 stabilized emulsions (Figure 6a). After 5 days of storage, propanol

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formation was in the order of lecithin > Tween 80 > SDS > quillaja saponin. With

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propanal formation, only lecithin-stabilized emulsion showed increased oxidation with

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light exposure (Figure 6b).

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Differences in the ability of the different emulsifiers to impact oxidation is most likely

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due to differences in their impact on hydroperoxide stability, since hydroperoxides must

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decompose to form propanal. In oil-in-water emulsions the major prooxidant that

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decomposes lipid hydroperoxides to propanal is iron.(34) The hydroperoxides in the

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Tween 80-stabilized emulsion could have been more stable because of the low charge of

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the emulsion droplet meaning that less iron is attracted to the emulsion droplet interface.

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Conversely, the lecithin-stabilized nanoemulsions are negatively charged and thus could


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attract iron, which could decompose the hydroperoxides into propanal thus explaining

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why lecithin-stabilized emulsion had low hydroperoxides and higher propanal levels than

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Tween 80-stabilized emulsions. SDS-stabilized nanoemulsions had intermediate

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oxidation rates even though it more negatively charged than the lecithin-stabilized

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nanoemulsions.

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nanoemulsions consistently had low oxidation rates even though it also produces

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negatively charged droplets.

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Impact of surfactant type on photosensitizer promoted oxidation in nanoemulsions

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We also examined lipid oxidation rates in nanoemulsions stabilized with the different

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surfactants in the presence of photosensitizersIn the presence of riboflavin, lipid

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hydroperoxides formation was greater for Tween 80 than SDS with the lecithin- and

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quillaja saponin-stabilized nanoemulsions having very low hydroperoxides levels after 5

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days of storage (Figure 7a). Headspace propanal formation after 5 days of storage were in

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the order of Tween 80 > lecithin > SDS > quillaja saponin (Figure 7b). In lecithin- and

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SDS-stabilized nanoemulsions hydroperoxides were low while propanal was high which

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was also seen in the absence of riboflavin (Figure 6a and 6b). This again could be due to

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iron being attracted to the negatively charged emulsion droplets where they would rapidly

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breakdown hydroperoxides into propanal thus explaining why hydroperoxides were low

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but propanal was high. Low hydroperoxide with high headspace aldehydes has also been

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observed in other negatively charged emulsion systems.(24) As with oxidation in the

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absence of riboflavin (Figure 6a and 6b), quillaja saponin-stabilized nanoemulsions had

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the lowest hydroperoxides and propanal.

It

is

unclear

why

this

occurred.


Quillaja

saponin-stabilized


314

One potential reason that the quillaja saponin-stabilized emulsions were chemically stable

315

with riboflavin promoted oxidation is that quillaja saponin absorbs light at wavelengths

316

similar to those that excite riboflavin (Figure 8) allowing it to inhibit reactive oxygen

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species generation by riboflavin which occurs at 355 nm.(14) Therefore, we also used rose

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bengal to generate, reactive oxygen species since Rose Bengal can generate singlet

319

oxygen at 530 nm(35) where quillaja saponin does not have strong absorption patterns

320

(Figure 7). In the presence of rose bengal, lipid hydroperoxides and headspace propanal

321

formation was similar for all four surfactants (Figure 9a and 9b). The suggests that the

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superior oxidative stability of quillaja saponin-stabilized emulsions in the presence of

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riboflavin compared to rose bengal was due to the ability of quillaja saponin to decrease

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riboflavin excitation and thus singlet oxygen generation.

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While quillaja saponin could inhibit riboflavin promoted oxidation by absorbing light at

326

similar wavelengths, the other surfactants could have done the same since they also

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absorb light at 355 nm with lecithin absorbing even more light than quillaja saponin.

328

However, nanoemulsions stabilized by quillaja saponin were more oxidatively stable than

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the other surfactant-stabilized emulsions in the absence of photosensitizers where

330

inhibition of riboflavin promoted oxidation would not be important. This suggests that

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quillaja saponin could inhibit lipid oxidation by other mechanisms such as free radical

332

scavenging. The ability of the surfactants to scavenge free radicals was evaluated by the

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ORAC assay. Tween 80 and SDS had the lowest radical scavenging capacity followed by

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lecithin (Table 2). Quillaja saponin’s free radical scavenging capacity (55.6 µM Trolox

335

Equivalences/µM surfactant) was 10 fold greater than lecithin and 55 fold greater than

336

SDS and Tween 80. Quillaja saponin is a saponin and saponins contain hydroxyl groups


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that could scavenge free radicals (Figure 2). Saponins from soy have previously been

338

found to scavenge free radicals and superoxide anion.(36) This could help explain why

339

quillaja saponin-stabilized nanoemulsions were more stable than the other surfactants in

340

both the presence and absence of riboflavin.

341

presence of riboflavin since photosensitization of riboflavin can also generate superoxide

342

anion which can increase the ability of iron to decompose hydroperoxides into free

343

radicals by reducing iron to the more reactive ferrous form.(19)

This would be especially true in the

344

In conclusion, ω-3 fortified nanoemulsion could be produced using stabilized using

345

both synthetic (Tween 80 and SDS) and natural (lecithin and quillaja saponin)

346

emulsifiers. The nanoemulsions prepared using Tween 80, quillaja saponin or SDS were

347

stable from pH 3 to 8, whereas those prepared using lecithin were unstable to droplet

348

aggregation and creaming below pH 5. The SDS- and quillaja saponin- stabilized

349

nanoemulsion were stable at NaCl concentrations from 0 to 500 nm and during thermal

350

processing for 30 min up to 90 oC, which was attributed to their strong electrostatic

351

charge which would inhibit droplet aggregation. Tween 80 and lecithin stabilized

352

nanoemulsions where less stable to NaCl and thermal processing. Quillaja saponin could

353

be an outstanding emulsifier for ω-3 ethyl esters nanoemulsions because it was physically

354

stable over a wide pH range (3-8), NaCl concentrations (≤ 500 mM) and thermal

355

processing conditions (up to 90°C). In addition, quillaja saponin consistently produced

356

the most oxidatively stable nanoemulsions in the presence and absence of

357

photosensitizers. This is likely due to the ability of quillaja saponin to scavenge free

358

radicals and in the case of riboflavin-promoted oxidation, absorb light in the range that

359

excites riboflavin.



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AUTHOR INFORMATION

361

Corresponding Author

362

*Phone: +90 (505) 272 0714. Fax: +90(422) 615 2060. E-mail: [email protected]

363

Notes

364

The authors declare no competing financial interest.

365

ACKNOWLEDGMENTS

366

The authors thank the Turkish Government for providing financial support for Sibel

367

Uluata. The authors also want to acknowledge TUBITAK 2219-A Program for providing

368

funding for Sibel Uluata.

369

This material was also partly based upon work supported by the Cooperative State

370

Research, Extension, Education Service, USDA, Massachusetts Agricultural Experiment

371

Station (Project No. 831) and USDA, NRI Grants (2011-03539, 2013-03795, 2011-

372

67021, and 2014-67021). This project was also partly funded by the Deanship of

373

Scientific Research (DSR), King Abdulaziz University, Jeddah, under grant numbers

374

330-130-1435-DSR, 299-130-1435-DSR, 87-130-35-HiCi. The authors, therefore,

375

acknowledge with thanks DSR technical and financial support.

376

ABBREVIATIONS USED

377

DHA: docosahexaenoic acid, EPA: eicosapentaenoic acid, SDS: Sodium dodecyl sulfate

378

ORAC: oxygen radical absorbance capacity


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(1) Flock, M. R.; Harris, W. S.; Kris-Etherton, P. M., Long-chain omega-3 fatty acids: time to establish a dietary reference intake. Nutrition Reviews 2013, 71, 692-707. (2) Dawn M. W.; Lashner B. A.; Lerner E.; DeMichele, S. J. S., D. L., The Effects of an Oral Supplement Enriched With Fish Oil, Prebiotics, and Antioxidants on Nutrition Status in Crohn’s Disease Patients. Nutrition in Clinical Practice 2011, 26. (3) Ellulu, M. S.; Khaza'ai, H.; Abed, Y.; Rahmat, A.; Ismail, P.; Ranneh, Y., Role of fish oil in human health and possible mechanism to reduce the inflammation. Inflammopharmacology 2015, 23, 79-89. (4) Pottel, L.; Lycke, M.; Boterberg, T.; Foubert, I.; Pottel, H.; Duprez, F.; Goethals, L.; Debruyne, P. R., Omega-3 fatty acids: physiology, biological sources and potential applications in supportive cancer care. Phytochemistry Reviews 2013, 13, 223-244. (5) Sun-Waterhouse, D., The development of fruit-based functional foods targeting the health and wellness market: a review. Inter. J. Food Sci. & Technol. 2011, 46, 899-920. (6) Coupland, J. N.; McClements D. J., Lipid oxidation in food emulsions. Trends in Food Sci. &Technol. 1996, 7, 83-90. (7) McClements, D. J., Edible nanoemulsions: fabrication, properties, and functional performance. Soft Matter 2011, 7, 2297. (8) McClements, D. J., Theoretical prediction of emulsion color. Adv. Coll. Inter. Sci. 2002, 97, 63-89. (9) Boon, C. S.; McClements, D. J.; Weiss, J.; Decker, E. A., Factors influencing the chemical stability of carotenoids in foods. Critical reviews in Food science and nutrition 2010, 50, 515-32. (10) Piorkowski, D. T.; McClements, D. J., Beverage emulsions: Recent developments in formulation, production, and applications. Food Hydrocolloids 2014, 42, 5-41. (11) McClements, D. J., Nanoemulsions versus microemulsions: terminology, differences, and similarities. Soft Matter 2012, 8, 1719. (12) Panya, A.; Laguerre, M.; Bayrasy, C.; Lecomte, J.; Villeneuve, P.; McClements, D. J.; Decker, E. A., An investigation of the versatile antioxidant mechanisms of action of rosmarinate alkyl esters in oil-in-water emulsions. J. Agric. Food Chem. 2012, 60, 2692700. (13) Choe, E.; Min, D.B., Mechanisms and Factors for Edible Oil Oxidation. Compre. Rev. Food Sci. Food Saftey 2006, 5, 169-186. (14) Baier, J.; Maisch, T.; Maier, M.; Engel, E.; Landthaler, M.; Baumler, W., Singlet oxygen generation by UVA light exposure of endogenous photosensitizers. Biophysic. J. 2006, 91, 1452-9. (15) DeRosa, M. C.; Cructchley, R. J., Photosensitized singlet oxygen and its applications. Coordination Chem. Reviews 2001, 233, 351-371. (16) Kim, H. J.; Min, D. B., Chemistry of lipid oxidation. In Food Lipids chemistry, Nutrition, and Biotechnology, third ed.; Casimir C. Akoh , D. B. M., Ed. Taylor and Francis Group: CRC press, 2008; p 299.



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(17) Min, D. B.; Boff, J. M., Chemistry and Reaction of Singlet Oxygen in Foods. Comph. Reviews Food Sci. Food Safeyt 2002, 1, 58-72. (18) Kim, T. S.; Decker, E. A.; Lee, J., Antioxidant capacities of α-tocopherol, trolox, ascorbic acid, and ascorbyl palmitate in riboflavin photosensitized oil-in-water emulsions. Food Chem. 2012, 133, 68-75. (19) Lee, J.; Decker, E. A., Effects of metal chelator, sodium azide, and superoxide dismutase on the oxidative stability in riboflavin-photosensitized oil-in-water emulsion systems. J. Agric. Food Chem. 2011, 59, 6271-6. (20) Huang, R.; Choe, E.; D.B. Min, Kinetics for Singlet Oxygen Formation by Riboflavin Photosensitization and the Reaction between Riboflavin and Singlet Oxygen. Food Chem. Toxicol. 2004, 69, 726-732.

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(21) Given, P. S., Encapsulation of Flavors in Emulsions for Beverages. Current Opinion in Colloid & Interface Science 2009, 14, 43-47. (22) Rao, J.; McClements, D. J., Food-grade microemulsions and nanoemulsions: Role of oil phase composition on formation and stability. Food Hydrocolloids 2012, 29, 326-334. (23) Fomuso, L. B.; Corredig, M.; Akoh, C.C., Effect of Emulsifier on Oxidation Properties of Fish Oil-Based Structured Lipid Emulsions. J. Agric. Food Chem 2002, 50, 2957-2961. (24) Mei, L.; McClements, D.J; Wu, J.; Decker E. A., Iron-catalyzed lipid oxidation in emulsion as affected by surfactant, pH and NaCl. Food Chemistry 1998, 61. (25) MCClements, D. J.; Decker, E.A, Lipid oxidation in oil-in-water Emulsions:Impact of molecular Environment on Chemical Reactions in Heterogeneous Food System. J. Food Sci. 2000, 65, 1270-1282. (26) Cho, Y. J.; McClements, D.J.; Decker, E. A., Ability of Surfactant Micelles To Alter the Physical Location and Reactivity of Iron in Oil-in-Water Emulsion. J Agric. Food Chem. 2002, 50, 5704-5710. (27) Hu, M.; McClements, D. J.; Decker, E.A., Impact of Whey Protein Emulsifiers on the Oxidative Stability of Salmon Oil-in-Water Emulsions. J. Agric. Food Chem 2003, 51, 1435-1439. (28) Walker, R.; Decker, E. A.; McClements, D. J., Development of food-grade nanoemulsions and emulsions for delivery of omega-3 fatty acids: opportunities and obstacles in the food industry. Food & Funct. 2015, 6, 42-55. (29) Zulueta, A.; Esteve, M. J.; Frígola, A., ORAC and TEAC assays comparison to measure the antioxidant capacity of food products. Food Chemistry 2009, 114, 310-316. (30) Uluata, S.; McClements, D. J.; Decker, E. A., How the multiple antioxidant properties of ascorbic acid affect lipid oxidation in oil-in-water emulsions. J. Agric. Food Chem. 2015, 63, 1819-24. (31) Panya, A. L., M.; Lecomte, J.; Villeneuve, P.; Weiss, J.; McClements, D. J.; Decker, E.A, Effects of chitosan and rosmarinate esters on the physical and oxidative stability of liposomes. J. Agric. Food Chem. 2010, 9, 5679−5684. (32) Yang, Y.; Leser, M. E.; Sher, A. A.; McClements, D. J., Formation and stability of emulsions using a natural small molecule surfactant: Quillaja saponin (Q-Naturale®). Food Hydrocolloids 2013, 30, 589-596. (33) Saberi, A. H.; Fang, Y.; McClements, D. J., Effect of Salts on Formation and Stability of Vitamin E-Enriched Mini-emulsions Produced by Spontaneous Emulsification. J. Agric. Food Chem. 2014, 62, 11246-11253.


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(34) Nuchi.C.D.; McClements, D. J.; Decker E. A., Impact of Tween 20 Hydroperoxides and Iron on the Oxidation of Methyl Linoleate and Salmon Oil Dispersions. J. Agric. Food Chem. 2001, 49, 4912-4916.

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(35) Bilski P.; Li, M.Y.; Ehrenshaft, M.; Daub, M. E.; Chignell, C. F., Symposium-inPrint Vitamin B6 (Pyridoxine) and Its Derivatives Are Efficient Singlet Oxygen Quenchers and Potential Fungal Antioxidants. Photochem. Photobiolo. 2000, 71, 129134. (36) Yoshiki, Y.; Kahara, T.; Okubo K.; Sakabe T.; and Yamasaki, T., Superoxide- and 1,1- Diphenyl-2-picrylhrazyl Radical-scavenging Activities of soyasaponin β g related to Gallic Acid. Biosci. Biotechnol. Biochem. 2001, 65, 2162-2165.

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491

Figure Captions

492

Figure 1. Appearance of 1 wt% fish oil ethyl ester nanoemulsions stabilized by Tween

493

80, SDS, lecithin or quillaja saponin.

494

Figure 2. Influence of storage pH on the mean particle diameter of 1 wt% fish oil ethyl

495

ester nanoemulsions stabilized by Tween 80, SDS, lecithin or quillaja saponin after 24 h.

496

Figure 3. Influence of salt concentration on mean particle diameter of 1 wt% fish oil

497

ethyl ester nanoemulsions stabilized by Tween 80, SDS, lecithin or quillaja saponin after

498

24 h.

499

Figure 4. Influence of temperature on mean particle diameter of 1 wt% fish oil ethyl ester

500

nanoemulsions stabilized by Tween 80, SDS, lecithin or quillaja saponin after 24 h.

501

Figure 5. Lipid hydroperoxide (a) and propanal (b) formation in 1 wt% fish oil ethyl

502

ester nanoemulsions stabilized by Tween 80, SDS, lecithin or at 37 oC. L: Light, D:

503

Dark.

504

Figure 6. Lipid hydroperoxide (a) and propanal (b) formation in of 1 wt% fish oil ethyl

505

ester nanoemulsions stabilized by Tween 80, SDS, Lecithin or quillaja saponin at 37 oC

506

in the presence of Riboflavin (Rb).

507

Figure 7. Absorbance spectra of Tween 80, SDS, lecithin or quillaja saponin, riboflavin

508

and rose bengal.

509

Figure 8. Lipid hydroperoxide (a) and propanal (b) formation of 1 wt% fish oil ethyl

510

ester nanoemulsions stabilized by Tween 80, SDS, lecithin or quillaja saponin at 37 oC in

511

the presence at Rose Bengal (Rose b).

512

Figure 9. Chemical Structure of quillaja saponin.(32)

513


Page 22 of 37

Page 23 of 37


514 515

Figure 1.

516

517

Quillaja saponin

SDS

Lecithin

518

519 520 521 522 523


Tween 80


524 525 526

Figure 2.

527 528 529 530 531 532 533 534


Page 24 of 37

Page 25 of 37


535 536

Figure 3.

Mean Particle diameter (nm)

150 Tween 80 SDS Lecithin Quillaja sapaonin

140 130 120 110 100 90 80 70 60 50 2

537

3

4

5

pH

538 539 540


6

7

8


Page 26 of 37

541 542

Figure 4.

Tween 80 SDS Lecithin Quillaja sapaonin


110

90

70

50

30 0 543

100

200

300

400

NaCl concentration(mM)

544 545 546


500

Page 27 of 37


547 548

Figure 5.

180 Tween 80 SDS Lecithin Quillaja saponin


160 140 120 100 80 60 40 20 30 549

40

50

60

70

Temperature (oC)

550 551 552


80

90


Page 28 of 37

553 554

Figure 6a.

Lipid Hydroperoxides (mmole/kg oil)

8

555

Tween 80-L SDS-L Lecithin-L Quillaja saponin-L Tween80-D SDS-D Lecithin-D Quillaja saponin-D

7 6 5 4 3 2 1 0 0

2

4

6

Oxidation time (day)

556 557 558


8

Page 29 of 37


559 560

Figure 6b.

400 Tween-L SDS-L Lecithin-L Quillaja saponin-L Tween80-D SDS-D Lecithin-D Quillaja saponin-D

Propanal(µmole/kg oil)

350 300 250 200 150 100 50 0 0 561

2

4

Oxidation time(day)

562 563 564


6

8


Page 30 of 37

565 566

Figure 7a.

7

Lipid Hydroperoxides (mmole/kg oil)

Tween 80-Rb SDS-Rb

6

Lecithin-Rb Quillaja saponin-Rb

5 4 3 2 1 0 0

567

2

4

6


568 569 570


8

Page 31 of 37


571 572 573

Figure 7b.

400 Tween80-Rb

350

SDS-Rb

Propanal (µmole/kg oil)

Lecithin-Rb

300

Quillaja saponin-Rb

250 200 150 100 50 0 0

574

2

4

oxidation time (day)

575 576


6

8


Page 32 of 37

577 578 579

Figure 8. 2

1.8

1.6

Absorbans (cm-1)

1.4

1.2 Tween 80 SDS 1

Lecithin Quillaja saponin

0.8

Riboflavin Rose Bengal

0.6

0.4

0.2

0 320

580

370

420

470

520

570

620

Wavelength (nm)

581 582


670

720

Page 33 of 37


583 584 585

Figure 9a.

Lipid hydroperoxides (mmole/kg oil)

8

Tween80-Rose b. SDS-Rose b. Lecithin-Rose b. Quillaja saponin-Rose b.

7 6 5 4 3 2 1 0 0

586

2

4

Oxidation time(day)

587 588


6

8


Page 34 of 37

589 590 591

Figure 9b.

400 Tween 80-Rose b.

350

SDS-Rose b.

Propanal (µmole/kg oil)

Lecithin-Rose b.

300

Quillaja saponin-Roseb

250 200 150 100 50 0 0

592

2

4


593 594


6

8

Page 35 of 37


595 596 597

Table 1. Mean particle diameter and droplet charge value of 1 wt% fish oil ethyl ester

598

nanoemulsions stabilized by Tween 80, SDS, lecithin or Quillaja saponin

599

Surfactant

Mean Particle Diameter (nm)

Droplet Charge (ζ-potential) (mV)

Tween 80

44.1 ± 0.8

-6.0 ± 1.2

SDS

81.6 ± 1.4

-62.5 ± 0.6

Lecithin

68.4 ± 1.3

-26.9 ± 1.6

Quillaja saponin

89.2 ± 0.2

-41.6 ± 2.6

600 601 602 603 604 605 606 607



608 609 610

Table 2. ORAC values of Tween 80, SDS, Lecithin and Quillaja saponin expressed as

611

µM TE/ µM surfactant.

Surfactant

ORACFL

Tween 80

1.1± 0.1c

SDS

0.5 ± 0.1d

Lecithin

5.4 ± 0.2b

Quillaja saponin

55.6 ± 1.2a

612

a

613

Letters indicate a significant difference (p≤ 0.05) between means.

A higher ORAC value represents greater free radical scavenging capacity.

614 615 616 617 618 619 620 621 622 623 624 625 626


Page 36 of 37

Page 37 of 37


627 628 629 630 631

TOC Graphic

632 633

Figure 1.

634 635

Quillaja saponin 636

SDS

Lecithin

637

638 639 640


Tween 80

Physical Stability, Autoxidation, and Photosensitized Oxidation of Ï-3

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