EMULSION COMPRISING ANTIOXIDANT PARTICLES

Abstract

The present invention relates to compositions comprising particles prepared from one or more biological materials and/or animal lipids and/or plant lipids that are capable of locating to an interface when combined with two or more immiscible liquids. Emulsions comprising the compositions comprising particles, wherein the emulsion has an internal phase dispersed in a continuous external phase and the particles are located at the interface of the external and the internal phase, methods of preparing such compositions and emulsions, the use of such compositions and emulsions and products containing the compositions and emulsions are also described.

Claims

1. A composition comprising particles prepared from one or more biological materials that are capable of locating to or at an interface when combined with two or more immiscible liquids.

2. The composition according to claim 1, wherein the particles comprise biological materials selected from the group consisting of blue-green algae, the Rutaceae family, the Malvaceae family, the Rubiaceae family, the Amaranthaceae family, the Poaceae family, the Zingiberaceae family, the Ginkgoaceae, the Araliaceae family, the Theaceae family, the Asteraceae family, the Oleaceae family, the Moringaceae family, the Bromeliaceae family, the Brassicaceae family, the Rosaceae family, the Sapindaceae family, the Lamiacea family, and mixtures thereof and/or animal lipids and/or plant lipids selected from the group consisting of milk fat, palm oil, palm kernel oil, coconut oil, cuphea oil, cocoa butter, shea butter, tripalmitin, palm stearin, waxes, fractionated oils, hydrogenated oils, and mixtures thereof.

3. (canceled)

4. The composition according to claim 2, wherein the particles prepared from animal lipids and/or plant lipids are solid at room temperature.

5. (canceled)

6. The composition according to claim 1, wherein the particles have a diameter from about 0.1 μm to about 100 μm.

7. The composition according to claim 1, wherein the particles comprise an antioxidant.

8. The composition according to claim 7, wherein the anti-oxidant is in or from a plant or microalgal extract rich in antioxidants.

9. The composition according to claim 8, wherein the plant or microalgal extract rich in antioxidants is a rosemary, sage, or green tea extract, raw material or extraction cake, a Dunaliella salina extract or oleoresin, a spirulina extract or extraction cake, or a spinach extract, raw material or extraction cake.

10. (canceled)

11. The composition according to claim 7, wherein the antioxidant is selected from the group consisting of tocopherols, tocotrienols, plastochromanols, phenolic diterpenes, flavonoids, phenolic acids and esters, stilbenes, carotenoids, essential oils and mixtures thereof, and/or synthetic antioxidants selected from the group consisting of butylated hydroxyanisole (BHA), butylated hydroxytoluene (BHT), tert-butyl-hydroxyquinone (TBHQ), propyl gallate (PG), ascorbyl palmitate and mixtures thereof.

12. (canceled)

13. An emulsion comprising a composition comprising particles as defined in claim 1, the emulsion comprising an internal phase dispersed in a continuous external phase, wherein particles are located at the interface of the external and the internal phase and at least one of the internal or external phase comprises an oxidisable compound.

14. The emulsion according to claim 13, wherein the oxidisable material comprises a lipid.

15. The emulsion according to claim 13 or 14, wherein the lipid has at least one carbon-carbon double bond in the fatty acyl chain and is selected from the group consisting of palmitoleic acid, oleic acid, myristoleic acid, linoleic acid, arachidonic acid, linolenic acid, eicosapentaenoic acid, docosahexaenoic acid, sunflower, soybean, canola, rapeseed, flaxseed, olive, peanut, corn, cottonseed, palm, fish oils, and combinations thereof.

16. The emulsion according to claim 13, wherein the emulsion is an oil-in-water emulsion.

17. (canceled)

18. (canceled)

19. (canceled)

20. A method for reducing or preventing oxidation and/or enhancing the oxidative stability of an emulsion comprising either: (ii) forming an emulsion comprising an internal phase dispersed in a continuous external phase and adding a composition comprising particles as defined in claim 1 to the emulsion; or (ii) forming an emulsion comprising an internal phase dispersed in a continuous external phase and a composition comprising particles as defined in claim 1 by mixing two or more immiscible liquids and the particles under conditions suitable for forming an emulsion; wherein at least one of the internal or external phase comprises an oxidisable material.

21. A method of prolonging the shelf-life of a beverage, a nutraceutical, a pharmaceutical or food product comprising an emulsion, wherein the method comprises either: (iii) forming an emulsion comprising an internal phase dispersed in a continuous external phase and adding a composition comprising particles as defined in claim 1 to the emulsion; or (iv) forming an emulsion comprising an internal phase dispersed in a continuous external phase and a composition comprising particles as defined in claim 1 by mixing two or more immiscible liquids and the particles under conditions suitable for forming an emulsion; wherein at least one of the internal or external phase comprises an oxidisable material.

22. (canceled)

23. (canceled)

24. (canceled)

25. The method according to claim 20, wherein the oxidisable material comprises a lipid.

26. The method according to claim 25, wherein the lipid has at least one carbon-carbon double bond in the fatty acyl chain and is selected from the group consisting of palmitoleic acid, oleic acid, myristoleic acid, linoleic acid, arachidonic acid, linolenic acid, eicosapentaenoic acid, docosahexaenoic acid, sunflower, soybean, canola, olive, peanut, corn, cottonseed, palm, fish oils, and combinations thereof.

27. The method according to claim 20, wherein the internal phase comprises oil and the external phase comprises water.

28. (canceled)

29. (canceled)

30. (canceled)

31. The method according to claim 20, wherein the particles reduce, delay and/or prevent the formation of oxidation products, secondary oxidation products and/or non-volatile secondary oxidation products.

32. The method according to claim 20, wherein the emulsion is a nutraceutical composition, dietary or food product for humans or animals, nutritional supplement, fragrance or flavouring, pharmaceutical or veterinary composition, oenological or cosmetic formulation or the emulsion is part of a nutraceutical composition, dietary or food product for humans or animals, nutritional supplement, fragrance or flavouring, pharmaceutical or veterinary composition, oenological or cosmetic formulation.

33. A nutraceutical composition, a dietary or food product for human or animals, nutritional supplements, a fragrance or flavouring, a pharmaceutical or veterinary composition, an oenological or cosmetic formulation comprising a composition as defined in claim 1.

34. A method of utilizing the composition as defined in claim 1 comprising adding said composition to a nutraceutical composition, a dietary or food product for humans or animals, a nutritional supplement, a fragrance or flavouring, a pharmaceutical or veterinary composition, an oenological or cosmetic formulation.

35. A method for the preparation of an emulsion, wherein the method comprises: mixing a composition as defined in claim 1 with either: (c) two or more immiscible liquids; or (d) a pre-prepared emulsion comprising an internal phase dispersed in a continuous external phase.

36. (canceled)

37. (canceled)

38. A kit for prolonging the shelf life of a beverage, a nutraceutical, a pharmaceutical or food product comprising an emulsion, wherein the emulsion comprises an internal phase dispersed in a continuous external phase, and at least one of the internal or external phase comprises an oxidisable material; the kit comprising particles as defined in claim 1.

39. The method according to claim 31, wherein the particles reduce, delay and/or prevent the formation of oxidation products including lipid hydroperoxides and conjugated diene hydroperoxides and/or secondary oxidation products including aldehyde, ketone, alcohol, and carboxylic acid volatile compounds and/or non-volatile secondary oxidation products including p-anisidine, epoxides, dimers and polymers.

40. A nutraceutical composition, a dietary or food product for human or animals, nutritional supplements, a fragrance or flavouring, a pharmaceutical or veterinary composition, an oenological or cosmetic formulation comprising a emulsion as defined in claim 13.

Description

BRIEF DESCRIPTION OF THE FIGURES

[0178] FIG. 1. Schematic representation of an oil-in-water emulsion stabilized by tripalmitin colloidal particles (A) or a conventional emulsifier where tripalmitin is solubilised in the oil phase (B).

[0179] FIG. 2. Lipid oxidation kinetics measured in conventional sodium caseinate-stabilized emulsion containing tripalmitin fat (circles) and Pickering emulsion stabilized by tripalmitin colloidal particles (squares). Both emulsions are incubated with 200 μM FeSO.sub.4/EDTA at 25° C. CD, conjugated dienes (left); p-AV, p-anisidine (right).

[0180] FIG. 3. Schematic representation of an oil-in-water emulsion stabilized by palm stearin colloidal particles (A) or stabilized by a conventional emulsifier where palm stearin is solubilised in the oil phase (B).

[0181] FIG. 4. Lipid oxidation kinetics measured in conventional sodium caseinate-stabilized emulsion containing palm stearin fat (circles) and Pickering emulsion stabilized by palm stearin colloidal particles (squares). Both emulsions are incubated with 200 μM FeSO.sub.4/EDTA at 25° C. CD, conjugated dienes (left); p-AV, p-anisidine (right).

[0182] FIG. 5. Characterization of the particle size distribution of some representative natural powders suspended in water at 1% (w/w). Matcha tea raw material (A), spinach leave raw material (B), spirulina extraction cake (C), pineapple fibers (D), and rosemary leave extraction cake (E). Non-micronized powder: solid line; micronized powder: dotted line.

[0183] FIG. 6. Scanning electron micrographs of the non-micronized and micronized (respectively) powders of matcha tea raw material (A and B), pineapple fibers (C and D), spinach raw material (E and F), rosemary leaves extraction cakes (G and H), spirulina extraction cakes (I and J), curcuma extract (K and L), and red radish extract (M and N).

[0184] FIG. 7. Particle size distribution of oil-in-water emulsions stabilized during three months at 4° C. by non-micronized natural powders or conventional emulsifiers. The tested powders are spinach (A), spirulina (B), matcha tea (C), pineapple fibers (D), while the conventional emulsifiers are Tween 60 at 1% (w/w) (E), egg yolk at 5% (w/w) (F). t.sub.0: solid line; t.sub.3: dotted line. All emulsions were prepared in a 50 mM acetate buffer pH 4.5 and contained 0.1 wt % potassium sorbate as antimicrobial.

[0185] FIG. 8. Particle size distribution of Pickering oil-in-water emulsions stabilized during one month at 4° C. by 5% wt non-micronized natural particles and added with 100 mM NaCl. Matcha tea (A), spinach leaves (B), and spirulina cakes (C). t.sub.0: solid line; t.sub.1: dotted line. All emulsions were prepared in a 50 mM acetate buffer pH 4.5.

[0186] FIG. 9. Particle size distribution of Pickering oil-in-water emulsions stabilized during one month at 4° C. by 5% wt non-micronized natural particles and added with 100 mM NaCl. Matcha tea (A), spinach leaves (B), spirulina cakes (C), and pineapple fibers (D). t.sub.0: solid line; t.sub.1: dotted line. All emulsions were prepared in a 50 mM phosphate buffer pH 7.0.

[0187] FIG. 10. Particle size distribution of Pickering oil-in-water emulsions stabilized during one month at 4° C. by 5% wt non-micronized pineapple fibers and added with 100 mM NaCl. t.sub.0: solid line; t.sub.1: dotted line. All emulsions were prepared in unbuffered ultrapure water of “Type 1” as defined by ISO3696 (for example, milliQ water).

[0188] FIG. 11. Particle size distribution of Pickering oil-in-water emulsions stabilized during one month at 4° C. by 5% wt non-micronized natural particles at acidic and neutral pH. Matcha tea at pH 4.5 (A) and pH 7.0 (B), and spirulina cakes at pH 4.5 (C) and pH 7.0 (D). t.sub.0: solid line; t.sub.1: dotted line. Emulsions at pH 4.5 were prepared in a 50 mM acetate buffer, while those at pH 7.0 were in a 50 mM phosphate buffer.

[0189] FIG. 12. Lipid oxidation kinetics measured in a conventional Tween 60-stabilized oil-in-water emulsion and in a Pickering oil-in-water emulsion stabilized by 5% (w/w) of a non-micronized spirulina cake powder. All emulsions are incubated at 25° C. for four months. All emulsions are prepared with a stripped sunflower oil and a 50 mM acetate buffer pH 4.5. They contain 0.1 wt % potassium sorbate as antimicrobial.

[0190] FIG. 13. Lipid oxidation kinetics measured in a conventional egg yolk-stabilized oil-in-water emulsion and in two Pickering oil-in-water emulsions stabilized by 5% (w/w) of a non-micronized matcha tea powder or 5% (w/w) of a non-micronized spinach leave powder. All emulsions are incubated at 25° C. for four months. All emulsions are prepared with a stripped sunflower oil and a 50 mM acetate buffer pH 4.5. They contain 0.1 wt % potassium sorbate as antimicrobial.

[0191] FIG. 14. Representative micrographs of Pickering water-in-oil emulsions (reverse emulsions) stabilized by 1% (w/w) non-micronized curcuma extract (A) and 2.5% (w/w) non-micronized rosemary leave extraction cakes (B).

[0192] FIG. 15. Surface-activity of the supernatants obtained after applying a washing procedure to the natural particles.

[0193] FIG. 16. Particle size distribution of oil-in-water emulsions stabilized during one week at 4° C. by 5% wt of the supernatant of washed non-micronized natural particles. Matcha tea (A), spinach leaves (B), spirulina cakes (C), and pineapple fibers (D). t.sub.0: solid line; t.sub.1: dotted line. All emulsions are prepared in a 50 mM acetate buffer pH 4.5.

[0194] FIG. 17. Particle size distribution of Pickering oil-in-water emulsions stabilized during four weeks (excepted for pineapple, 1 week) at 4° C. by 5% wt of washed non-micronized natural particles. Matcha tea (A), spinach leaves (B), spirulina cakes (C), and pineapple fibers (D). t.sub.0: solid line; t.sub.4: dotted line. All emulsions are prepared in a 50 mM acetate buffer pH 4.5.

[0195] FIG. 18. Schematic representation of two types of colloidal particle-stabilized stripped sunflower oil-in-water emulsions (Pickering emulsions). Emulsion composition is identical, but α-tocopherol is incorporated in the colloidal particles (the emulsion of the invention) (A) or in the liquid PUFA oil droplets (control emulsion) (B). Conjugated diene hydroperoxide (CD-LOOH) concentration (C), p-anisidine value (p-AV) (D) and α-tocopherol degradation (E) in both oil-in-water emulsions incubated at 25° C. with 200 μM FeSO.sub.4/EDTA. Averaged values±standard deviations result from independent triplicates. Symbols: the emulsion of the invention (A); square grey symbols: the control emulsion (B) black circles. Both Pickering emulsions are stabilized by tripalmitin colloidal particles.

[0196] FIG. 19. Confocal laser scanning microscopy images of the Pickering stripped sunflower oil-in-water emulsion of the invention (A) and of the control emulsion (B) with 25-NBD-cholesterol (fluorescent analogue of α-tocopherol) initially added in tripalmitin colloidal particles (A) or within the droplets (B), taken at different time points. Polarized light microscopy images of the Pickering emulsion of the invention produced with colloid mill homogenization at t.sub.0 and t.sub.72 h (C and D, respectively). In panels A and B, the scale bar represents 10 μm. Both Pickering emulsions are stabilized by tripalmitin colloidal particles.

[0197] FIG. 20. DSC melting and crystallization thermograms of the Pickering stripped sunflower oil-in-water emulsion of the invention stabilized by tripalmitin colloidal particles containing α-tocopherol at t.sub.0 and t.sub.336 (C).

[0198] FIG. 21. Schematic representation of two types of colloidal particle-stabilized stripped sunflower oil-in-water emulsions (Pickering emulsions). Emulsion composition is identical, but carnosic acid is incorporated in the tripalmitin colloidal particles (the emulsion of the invention) (A) or in the liquid PUFA oil droplets (control emulsion) (B). Conjugated diene hydroperoxide (CD-LOOH) concentration (C) and p-anisidine value (p-AV) (D) in both oil-in-water emulsions incubated at 25° C. with 200 μM FeSO.sub.4/EDTA. Averaged values±standard deviations result from independent triplicates. Symbols: the emulsion of the invention (A); square grey symbols: the control emulsion (B) black circles.

[0199] FIG. 22. Schematic illustration of two types of conventional sodium caseinate-stabilized stripped sunflower oil-in-water emulsions comprising a suspension of tripalmitin colloidal particles in their aqueous phase. Emulsion and suspension composition is identical, but antioxidant is located either in the suspended tripalmitin colloidal particles (A) or in the core of the oil droplets (B). Total antioxidant concentration is similar in both systems. Conjugated diene hydroperoxide (CD-LOOH) content (C), p-anisidine value (p-AV) (D), and α-tocopherol recovery (E) of the two types of emulsions with α-tocopherol in the suspended colloidal particles (black circles) or in the liquid oil droplets (grey squares), incubated with 200 μM FeSO.sub.4/EDTA at 25° C. Averaged values+/−standard deviation result from independent triplicates (C, D, and E).

[0200] FIG. 23. Confocal laser scanning microscopy images of the conventional stripped sunflower oil-in-water emulsion with added tripalmitin colloidal particles with 25-NBD-cholesterol (fluorescent analogue of α-tocopherol) initially added in the palmitin colloidal particles (A) or in the droplets (B), taken at different time points. In panels A and B, the scale bar represents 10 μm.

[0201] FIG. 24. Schematic illustration of Pickering stripped sunflower oil-in-water emulsions comprising an interfacially-adsorbed population of antioxidant-free tripalmitin colloidal particles and an aqueous phase-suspended population of tripalmitin colloidal particles containing α-tocopherol (A) or not (B). Emulsion and suspension composition is identical, but antioxidant is located either in the suspended colloidal particles (A) or in the core of the oil droplets (B). Total antioxidant concentration is similar in both systems. Conjugated diene hydroperoxide (CD-LOOH) content (C), p-anisidine value (p-AV) (D), and a-tocopherol recovery (E) of the two types of emulsions with α-tocopherol in the suspended colloidal particles (black circles) or in the liquid oil droplets (grey squares), incubated with 200 μM FeSO.sub.4/EDTA at 25° C. Averaged values+/−standard deviation result from independent triplicates (C, D, and E).

[0202] FIG. 25. Confocal laser scanning microscopy images taken at different time points of the Pickering stripped sunflower oil-in-water emulsions comprising an interfacially-adsorbed population of antioxidant-free palmitin colloidal particles and an aqueous phase-suspended population of tripalmitin colloidal particles containing 25-NBD-cholesterol (fluorescent analogue of α-tocopherol, A) or not (B). In this latter case, the fluorescent analogue is initially located in the oil droplets. In panels A and B, the scale bar represents 10 μm.

[0203] FIG. 26. Conjugated diene hydroperoxide (CD-LOOH) concentration (A), p-anisidine value (p-AV) (B), and α-tocopherol degradation (C) during incubation of the Pickering stripped sunflower oil-in-water emulsion of the invention containing 90 (black circles), 45 (triangles) or 22.5 (diamonds) ppm of α-tocopherol in the tripalmitin colloidal particles, and a control Pickering oil-in-water emulsion containing 90 ppm of α-tocopherol in the oil droplets (grey squares).

[0204] FIG. 27. Characterization of tripalmitin colloidal particles with or without α-tocopherol. Particle size distribution (A), DSC melting and crystallization thermograms (B), and TEM image of tripalmitin colloidal particles with tocopherol (C).

[0205] FIG. 28. Characterization of Pickering stripped sunflower oil-in-water emulsions with a-tocopherol either in the tripalmitin colloidal particles (the emulsion of the invention) or in the core of the oil droplets (control emulsion). Droplet size distribution (A), DSC melting and crystallization thermogram (B) and TEM image (C) of the emulsion of the invention.

[0206] FIG. 29. Stability of α-tocopherol during incubation of Pickering oil-in-water emulsions containing medium chain triglycerides with α-tocopherol either in the tripalmitin colloidal particles (black circles) or in the core of the oil droplets (grey squares).

[0207] FIG. 30. DSC melting and crystallization thermograms of a conventional sodium caseinate-stabilized stripped sunflower oil-in-water emulsion comprising tripalmitin colloidal particles in the aqueous phase (solid line), and a tripalmitin colloidal particle dispersion (dashed line).

[0208] FIG. 31. Schematic representation of two types of colloidal particle-stabilized stripped sunflower oil-in-water emulsions (Pickering emulsions). Emulsion composition is identical, but α-tocopherol is incorporated in the particles (the emulsion of the invention) (A) or in the liquid PUFA oil droplets (control emulsion) (B). Conjugated diene hydroperoxide (CD-LOOH) concentration (C), p-anisidine value (p-AV) (D) and α-tocopherol degradation (E) in both oil-in-water emulsions incubated at 25° C. with 200 μM FeSO.sub.4/EDTA. Averaged values±standard deviations result from independent triplicates. Symbols: the emulsion of the invention (A); grey squares: the control emulsion (B) black circles. Both Pickering emulsions are stabilized by tripalmitin (80%) colloidal particles containing 20% (w/w) of liquid tricaprylin.

[0209] FIG. 32. Confocal laser scanning microscopy images of the Pickering stripped sunflower oil-in-water emulsion of the invention (A) and of the control emulsion (B) with 25-NBD-cholesterol (fluorescent analogue of α-tocopherol) initially added in colloidal particles (A) or within the droplets (B), taken at different time points. Polarized light microscopy images of the Pickering emulsion of the invention produced with colloid mill homogenization at t.sub.0 and t.sub.72 h (C and D, respectively). In panels A and B, the scale bar represents 10 μm. Both Pickering emulsions are stabilized by tripalmitin colloidal particles.

[0210] FIG. 33. Schematic representation of two types of colloidal particle-stabilized non stripped flaxseed oil-in-water emulsions (Pickering emulsions). Emulsion composition is identical, but α-tocopherol is incorporated in the particles (the emulsion of the invention) (A) or in the liquid PUFA oil droplets (control emulsion) (B). Conjugated diene hydroperoxide (CD-LOOH) concentration (C) in both oil-in-water emulsions incubated at 25° C. with 200 μM FeSO.sub.4/EDTA. Averaged values±standard deviations result from independent triplicates. Symbols: the emulsion of the invention (A); grey squares: the control emulsion (B) black circles. Both Pickering emulsions are stabilized by tripalmitin colloidal particles.

EXAMPLES

[0211] The present invention will be further described by reference to the following, non-limiting examples.

Material and Methods

[0212] 1) Materials

[0213] Tripalmitin (#T8127, purity >99%), sodium phosphate monobasic (#S9638), sodium phosphate dibasic (#S9763), sodium chloride (#S7653), iron(II) sulfate heptahydrate (#F8633), ethylenediaminetetraacetic acid disodium salt dihydrate (#E6635), para-anisidine (#A88255), and acetic acid (#45726) were purchased from Sigma-Aldrich. N-Hexane (#808023502) was obtained from Actu-All Chemicals (Oss, the Netherlands). 2-Propanol was purchased from Merck (Darmstadt, Germany). Sodium caseinate was supplied by DMV International (#41610, spray dried, protein content 91.0%). Sunflower oil was obtained from a local supermarket, and was stripped with alumina powder (MP EcoChrome™ ALUMINA N, Activity: Super I, Biomedicals) to remove impurities and tocopherols. Palm stearin (palmitic acid, 82%; oleic acid, 9%; stearic acid, 5%) was supplied by ADM (Saint Laurent Bangy, France). Ultrapure water (18.2 MΩ) was used for all experiments, and was prepared using a Milli-Q system (Millipore Corporation, Billerica, Mass., USA). All other chemicals or solvents were of analytical grade.

[0214] 2) Purification of Tripalmitin

[0215] Tripalmitin was purified by three recrystallization steps using ethanol. Briefly, tripalmitin was dissolved in ethanol at 60-70° C. while stirring for 15 min and left to cool down to room temperature to allow recrystallization, after which ethanol was removed, which was repeated two more times.

[0216] 3) Preparation of Colloidal Lipid Particles (CLPs)

[0217] An aqueous phase containing sodium caseinate in phosphate buffer (10 mM, pH 7.0) was heated in a water bath and added to a melted fat phase (tripalmitin, palm stearin or tricaprylin).

[0218] When the particles contained tocopherol, 100 μL α-tocopherol prepared in methanol (200 mg mL.sup.−1) was added at this stage. Final α-tocopherol concentrations were 4 mg mg.sup.−1 of fat.

[0219] A coarse emulsion was then prepared by high speed stirring.

[0220] The coarse emulsion was then homogenized at high pressure and temperature then left to cool down, allowing for the lipid phase to crystallize.

[0221] 4) A General Procedure for the Preparation of O/W Emulsions for Studying the Antioxidative Effect of Particles Prepared from One or More Biological Materials

[0222] Two types of oil in water emulsions were prepared: one Pickering emulsion, stabilized by colloidal lipid particles (CLP) (tripalmitin or palm stearin) as the one or more biological materials (FIGS. 1A and 3A); and a conventional sodium caseinate-stabilized oil-in-water emulsion containing the same HMP fat (tripalmitin or palm stearin) in its oil interior (FIGS. 1B and 3B). Both emulsions contained the same amount of HMP fat but differed in their structural organization.

[0223] For the conventional sodium caseinate-stabilized emulsion containing HMP fat, stripped sunflower oil was mixed with tripalmitin, phosphate buffer (10 mM, pH 7.0) and sodium caseinate in phosphate buffer (10 mM, pH 7.0) at elevated temperature.

[0224] For the CLP-stabilized Pickering emulsion, stripped sunflower oil was mixed with phosphate buffer (10 mM, pH 7.0) and a particle dispersion.

[0225] The O/W emulsions were processed either by high pressure homogenization or colloid mill homogenization.

[0226] Coarse emulsions were prepared by high speed stirring. The obtained emulsions were then either homogenized at high pressure or processed through a colloid mill.

[0227] 5) A General Procedure for the Preparation of O/W Emulsions for Studying the Antioxidative Effect of Particles Prepared from One or More Biological Materials Filled with an Antioxidant

[0228] Two types of oil in water Pickering emulsions were prepared; one with α-tocopherol in the particles, and one with α-tocopherol in the liquid sunflower oil droplets (FIGS. 18, 21, 31, and 33).

[0229] In the former case, sunflower oil, preliminary stripped from surface-active impurities, was mixed with phosphate buffer (10 mM, pH 7.0) and a particle dispersion (with α-tocopherol in the particles).

[0230] In the latter case, components were mixed in the same proportions, but the particles did not contain α-tocopherol, whereas the sunflower oil was added with 100 μL α-tocopherol prepared in methanol (200 mg mL.sup.−1), before homogenization.

[0231] The mixtures were processed by high speed stirring. The obtained emulsions were then homogenized at high pressure and stored at cold temperature.

[0232] 6) Extraction and Analysis of α-Tocopherol

[0233] α-Tocopherol was extracted from CLPs dispersions or emulsions. First, 4 mL chloroform, 3 mL methanol and 1 mL saturated sodium chloride solution were added to 2 mL of CLP dispersion or emulsion in a 15-mL polypropylene centrifugation tube, which were vortexed followed by centrifugation at 3000×g for 10 minutes. The clear chloroform phase was then collected by cautiously boring a hole in the bottom of the centrifugation tube.

[0234] Extracts were analysed on a UltiMate 3000 liquid chromatography system (Thermo Scientific, Sunnyvale, Calif., USA) using a C30 reversed phase column, 3 μm, 150×4.6 mm (YMC, Dinslaken, Germany). Extracts were eluted at 1 mL min′ at 30° C. using a mobile phase with a linear gradient going from 81% methanol, 14% methyl t-butyl ether (MTBE) and 4% Milli-Q water to 74% methanol, 22% methyl t-butyl ether and 4% Milli-Q water in 8 minutes, and going back to its initial composition in 2 minutes. α-Tocopherol was detected with a UV-VIS detector at 292 nm (Dionex™ UltiMate™ 3000 Variable Wavelength Detector), and contents were calculated using a calibration curve that was linear in the range from 5 μg mL.sup.−1 to 5000 μg mL.sup.−1. The recovery (Rec %) of α-tocopherol in CLPs was calculated as:

[00001]$\mathrm{Rec}\%=100\frac{C_{\mathrm{ex}}}{C_{\mathrm{in}}}$

where C.sub.ex is the content of extracted α-tocopherol and C.sub.in the content of initially added a-tocopherol.

[0235] 7) Lipid Oxidation Experiments

[0236] A catalyst consisting of an equimolar mixture of FeSO.sub.4 and EDTA was prepared by separately dissolving FeSO.sub.4 and EDTA (12 mM) in ultrapure water. Equivalent volumes of each solution were mixed, and the iron-EDTA complex was allowed to form under moderate stirring for 1 h in the dark (Berton, Ropers, Viau, & Genot, 2011). Aliquots of emulsion (2 g) were distributed in a 15-mL polypropylene centrifugation tube. The catalyst (100 μL) was added to the emulsions to obtain a final concentration of 200 μM of both iron and EDTA. The tubes were rotated in the dark at 2 rpm at 25° C. for 0 to 72 h.

Formation of Conjugated Diene Hydroperoxides (CD-LOOH).

[0237] Quantification of CD-LOOH, which are primary lipid oxidation products, was adapted from Corongiu & Banni (1994). In short, the incubated emulsions were diluted 4000-fold in 2-propanol in multiple steps. The final solutions were centrifuged at 20238×g for 1 minute (Centrifuge 5424, Eppendorf Hamburg, Germany), and the absorbance of the supernatant was measured at 233 nm with a UV-visible spectrophotometer (DU 720 Beckman Coulter, Brea, Calif., USA). The reference cell contained 2-propanol and phosphate buffer (10 mM, pH 7.0) in the same proportions as in the final dilution of the samples. Results were expressed in mmol of equivalent hydroperoxides per kg of oil (mmol eq HP kg.sup.−1 oil) with 27000 M.sup.−1 cm.sup.−1 as the molar extinction coefficient of CD at 233 nm.

Formation of Total Aldehydes.

[0238] The para-anisidine value (p-AV), a measure of total aldehydes, was used to assess the formation of secondary lipid oxidation products (AOCS, 1998). In short, 1 mL saturated sodium chloride solution and 5 mL hexane/isopropanol (1/1, v/v) were added per aliquot of incubated emulsion (2.1 mL). Mixtures were vortexed followed by centrifugation at 2000×g for 8 minutes at 4° C. The upper hexane layer (>2 mL) was collected and placed on ice for 3 minutes, followed by centrifugation at 20238×g for 1 minute. The absorbance of the supernatant was measured at 350 nm with pure hexane as a blank (Ab). In a centrifugation vial, 1 mL of the supernatant was mixed with 0.2 mL 2.5 g/L para-anisidine in acetic acid solution. After exactly 10 min, the absorbance was measured at 350 nm using 1 mL pure hexane mixed with 0.2 mL 2.5 g/L para-anisidine in acetic acid solution, incubated for 10 min, as a blank (As). The para-anisidine value (pAV, arbitrary units) was calculated as follows:

[00002]$\mathrm{pAV}=\frac{\left(1.2\mathrm{As}-\mathrm{Ab}\right)}{m}$

[0239] Where m is the concentration of oil (g/mL).

Example 1. Oxidative Stability of a Conventional Sodium Caseinate-Stabilized Oil in Water Emulsion Containing Tripalmitin Compared to an Emulsion of the Invention Comprising Tripalmitin Colloidal Particles Prepared as Detailed in the Material and Methods Section

[0240] The oxidative stability of a Pickering emulsion stabilized by tripalmitin colloidal particles (PTP) has been evaluated by both conjugated dienes (primary oxidation products) and p-anisidine (secondary oxidation products) in comparison to a conventional sodium caseinate-stabilized emulsion containing HMP fat in the same amount (FIG. 1). Lipid oxidation in both emulsions was accelerated by 200 μM FeSO.sub.4/EDTA. The data obtained showed that tripalmitin colloidal particles exert a protective effect on the corresponding Pickering emulsion as both CD-LOOH and p-AV raised more slowly compared to the same emulsion (conventional) wherein tripalmitin is dissolved in the interior of the oil droplet (FIG. 2).

Example 2. Oxidative Stability of a Conventional Sodium Caseinate-Stabilized Oil in Water Emulsion Containing Palm Stearin Compared to an Emulsion of the Invention Comprising Palm Stearin Colloidal Particles Prepared as Detailed in the Material and Methods Section

[0241] The same experiment as in Example 1 was repeated using palm stearin instead of tripalmitin. A Pickering emulsion stabilized by colloidal particles formed by palm stearin (PPS) was evaluated in comparison to a conventional sodium caseinate-stabilized emulsion containing HMP fat in the same amount (FIG. 3). Lipid oxidation in both emulsions was accelerated by 200 μM FeSO.sub.4/EDTA. The effect previously seen with tripalmitin was exacerbated with palm stearin. The colloidal particles in this example exerted a huge antioxidative effect on the corresponding Pickering emulsion. This has been demonstrated on both primary (conjugated dienes) and secondary (p-anisidine) oxidation products (FIG. 4).

Example 3. Characterization of the Particle Size Distribution of Particles Prepared from One or More Biological Materials in Suspension where the Biological Material is Obtained from a Photosynthetic Organism

[0242] The characterization of the particle size distribution of some representative natural powders suspended in water at 1% (w/w) was performed using static light scattering (Malvern Mastersizer 3000, Malvern Instruments Ltd., Malvern, Worcestershire, UK) with a refractive index particle of 1.45 and an adsorption index of 0.01 (FIG. 5). Both micronized and non-micronized powders (matcha tea raw material, spinach leaves raw material, spirulina extraction cake, pineapple fibers, and rosemary leave extraction cake) were tested.

[0243] Interestingly, no particle size below 0.2 μm was measured demonstrating that the particles used were not nanoscale.

[0244] Particles from pineapple and spinach leave powders were found to possess a higher particle size than matcha tea and spirulina cake. The non-micronized spinach leaves particles contained particles of various diameters (i.e. a polydisperse distribution) with a main peak at 200 μm, whereas the micronized particles of the same material contained particles of uniform size (i.e. had a monodisperse distribution) with an average particle size of 8 μM. These results also show that micronization not only has a significant impact on the reduction of particle size, but also on the size distribution.

[0245] Matcha tea powder and spinach leave particles were polydispersed before micronization, but monodispersed after processing. Spirulina cake became more polydispersed once micronized, whereas pineapple kept a monodisperse distribution. The rosemary cake powder in both micronized and non-micronized form was polydisperse in size, appearing as big and small particles. Nevertheless, unlike the other materials, the particle size distribution was not significantly affected by micronization.

Example 4. Chemical Characterization of Particles Prepared from One or More Biological Materials where the Biological Material is Obtained from a Photosynthetic Organism

[0246] A chemical characterization of a representative set of micronized or non-micronized particles has been done and is presented in Tables 1 and 2.

TABLE-US-00001 TABLE 1 Composition table of some representative particles. Polyphenol content values with asterisks were determined by HPLC, while the others were from the Folin-Ciocalteu method. Free Total Free Total Samples Maltodextrins sugars Sugars glucose glucose Starch Ash Polyphenols Micronized ND ND 0.17 ND ND ND 1.36 92.91* curcumin Non-micronized ND ND 0.19 ND ND ND ND 92.75* curcumin Micronized ND 1.62 4.89 ND ND ND 1.50 11.27 rosemary cake Non-micronized ND 2.2  4.17 0.25 ND ND 1.58 10.95 rosemary Micronized 42.36 ND 84.61 ND 78.23  27.26 ND 1.33 red radish Non-micronized 44.06 0.52 94.01 0.52 77.1  24.10 ND 1.62 red radish Micronized ND 0.2  5.57 0.12 ND ND 17.70 0.15 spirulina Non-micronized ND 0.2  5.79 0.2  ND ND 17.79 0.36 spirulina Micronized ND 3.38 16.61 0.15 ND ND 4.72 14.54 matcha tea Non-micronized ND 4.95 15.7 0.73 ND ND 4.70 21 matcha tea Micronized ND 1.16 33.0 0.52 ND ND 12 0.47 pineapple Non-micronized ND 0.88 31.06 ND ND ND 1.09 1.45 pineapple Micronized ND 4.82 17.21 1.16 4.95  3.38 14.19 1.17 spinach Non-micronized ND 5.22 17.75 0.79 5.14  3.87 15 1.11 spinach

TABLE-US-00002 TABLE 2 Follow-up composition table of some representative particles. Neutral Total detergent Samples Proteins nitrogen Cellulose Fibres Fibres Lignin Hemicellulose Micronized <0.08 <0.5 <2.00 2.20 <0.50 <0.50 2.20 curcumin Non-micronized 1.2 0.18 14.6 34.2 17.7 16.8 16.5 rosemary Micronized red 0.50 0.08 <2.00 3.40 1.20 <0.50 2.20 radish Non-micronized 0.90 0.14 <2.00 3.80 1.30 <0.50 2.50 red radish Micronized 57.8 9.25 <2.00 5.50 3.30 <0.50 2.20 spirulina Non-micronized 58.10 9.30 <2.00 1.20 0.60 <0.50 <0.60 spirulina Micronized 22.70 3.64 2.80 5.10 2.90 1.20 2.20 matcha Non-micronized 22.00 3.52 4.60 27.60 16.40 8.50 11.20 matcha Micronized 1.80 0.28 17.90 26.20 13.20 7.10 13.00 pineapple Non-micronized 1.60 0.26 28.60 76.20 35.60 9.60 40.60 pineapple Micronized 28.85 4.62 6.95 15.6 5.45 2.75 10.05 spinach not micronized 28.00 4.48 6.70 15.10 8.40 1.30 6.70 Spinach

Example 5. Morphological Characterization of Particles Prepared from One or More Biological Materials where the Biological Material is Obtained from a Photosynthetic Organism in their Dry Form

[0247] The particles dried form microstructure was accessed using scanning electron microscopy (SEM). The non-micronized curcuma particles had a polyhedral shape whereas the micronized sample had an irregular shape (FIG. 6). The non-micronized red radish and spirulina cake particles had initially a spherical shape, but after processing, both micronized samples were irregular. Therefore, the results showed that for these particles, the microstructure was broken down by ultrasonification (i.e. micronization).

[0248] For matcha tea powder, pineapple, rosemary cake and spinach leaves particles shape did not seem to be affected by micronization as both non-micronized and micronized particles presented an irregular structure before and after processing. Moreover, the matcha tea powder, spinach leaves and rosemary cake particles presented high porosity, whereas the pineapple particles did not.

Example 6. Characterization of the Physical Stability of the Pickering Oil-in-Water Emulsions of the Invention Stabilized by Particles Prepared from One or More Biological Materials where the Biological Material is Obtained from a Photosynthetic Organism

[0249] In this example, the emulsion forming and stabilizing ability of the natural particles of the invention was assessed through the particle size distribution of the corresponding oil-in-water emulsions. FIG. 7 shows that spinach leaves (A), spirulina cake (B), matcha tea (C), and pineapple fibers (D), all in their non-micronized form were able, when added at 5% w/w, to form and stabilize Pickering oil-in-water emulsions over three months at 4° C.

[0250] These emulsions were formed and stabilized in an emulsifier-free medium and were compared to two conventional oil-in-water emulsions stabilized by Tween 60 at 1% (w/w) (FIG. 7E) or egg yolk at 5% (w/w) (FIG. 7F). All emulsions were prepared in a 50 mM acetate buffer pH 4.5 and contained 0.1 wt % potassium sorbate as antimicrobial.

Example 7. Characterization of the Physical Stability of the Pickering Oil-in-Water Emulsions of the Invention Stabilized by Particles Prepared from One or More Biological Materials where the Biological Material is Obtained from a Photosynthetic Organism when NaCl is Added

[0251] In this example, the emulsion forming and stabilizing ability of particles prepared from one or more biological materials where the biological material is obtained from a photosynthetic organism were assessed in the presence of NaCl through the particle size distribution of the corresponding oil-in-water emulsions.

[0252] FIG. 8 shows that matcha tea (A), spinach leaves (B), and spirulina cakes (C), all in their non-micronized form, were able, when added at 5% w/w, to form and stabilize (during one month at 4° C.) Pickering oil-in-water emulsions prepared in a 50 mM acetate buffer pH 4.5 in presence of a substantial level of salt.

[0253] FIG. 9 shows similar results for non-micronized matcha tea (A), spinach leaves (B), spirulina cakes (C), and pineapple fibers (D) when the exact same emulsions were prepared in a 50 mM phosphate buffer pH 7.0 instead of an acetate buffer.

[0254] Finally, FIG. 10, shows similar results for non-micronized pineapple fibers when the exact same emulsion was prepared in unbuffered ultrapure water of “Type 1” as defined by ISO3696 (for example, milliQ water).

[0255] This series of data clearly indicates that the Pickering oil-in-water emulsions of the invention can be formed and stabilized for a significant amount of time in presence of salt which is known for having in some cases disturbing effect on the physical stability of oil-in-water emulsions.

Example 8. Characterization of the Physical Stability of the Pickering Oil-in-Water Emulsions of the Invention Stabilized by Particles Prepared from One or More Biological Materials where the Biological Material is Obtained from a Photosynthetic Organism at Neutral and Acidic pH

[0256] Here, the emulsion forming and stabilizing ability of particles prepared from one or more biological materials where the biological material is obtained from a photosynthetic organism was assessed at different pH through the particle size distribution of the corresponding oil-in-water emulsions. FIG. 11 shows that matcha tea powder at pH 4.5 (A) and pH 7.0 (B), as well as spirulina cake powder at pH 4.5 (C) and pH 7.0 (D), all in their non-micronized form, were able, when added at 5% w/w, to form and stabilize (during one month at 4° C.) Pickering oil-in-water emulsions in presence of a substantial level of salt. Emulsions at pH 4.5 were prepared in a 50 mM acetate buffer, while those at pH 7.0 were in a 50 mM phosphate buffer.

Example 9. Oxidative Stability of a Conventional Tween 60-Stabilized Oil in Water Emulsion Compared to a Pickering Oil-in-Water Emulsion of the Invention Stabilized by 5% (w/w) of a Non-Micronized Spirulina Cake Powder

[0257] The oxidative stability of a conventional Tween 60-stabilized oil in water emulsion has been evaluated through the level of conjugated dienes (conjugated E,Z-Ln-OOH, primary oxidation products), lipid hydroperoxides (LOOHs, primary oxidation products) and aldehydes (secondary oxidation products) in comparison to a Pickering oil-in-water emulsion (emulsion of the invention) stabilized by 5% (w/w) of a non-micronized spirulina cake powder (FIG. 12). Lipid oxidation in both emulsions was natural (i.e. non-accelerated by oxidation catalyst(s) other than those naturally present in the systems). After four months of incubation at 25° C., data shows that spirulina cake powder exerts a surprising protective effect on the corresponding Pickering oil-in-water emulsion as all oxidation markers raised more slowly compared to the conventional emulsion stabilized by Tween 60 (a standard emulsifier used in industry). Thus, the particles of spirulina cakes act as natural antioxidant colloids or particles.

Example 10. Oxidative Stability of a Conventional Egg Yolk-Stabilized Oil in Water Emulsion Compared to Two Pickering Oil-in-Water Emulsions of the Invention Stabilized by 5% (w/w) of a Non-Micronized Matcha Tea Powder or 5% (w/w) of a Non-Micronized Spinach Leave

[0258] The oxidative stability of a conventional egg yolk-stabilized oil in water emulsion has been evaluated through the level of conjugated dienes (conjugated E,Z-Ln-OOH, primary oxidation products), lipid hydroperoxides (LOOHs, primary oxidation products) and aldehydes (secondary oxidation products) in comparison to two Pickering oil-in-water emulsions (emulsions of the invention) stabilized by 5% (w/w) of non-micronized matcha tea powder or non-micronized spinach leaves (FIG. 13). Lipid oxidation in both emulsions was natural (i.e. non-accelerated by oxidation catalyst(s) other than those naturally present in the systems). After four months of incubation at 25° C., data shows that both natural powders exert a surprising protective effect on the corresponding Pickering oil-in-water emulsions as all oxidation markers raised more slowly compared to the conventional emulsion stabilized by egg yolk (a standard emulsifier used in industry). Thus, the particles of matcha tea and those of spinach leaves act as natural antioxidant colloids or particles.

Example 11. Characterization of the Ability of Particles Prepared from One or More Biological Materials where the Biological Material is Obtained from a Photosynthetic Organism to Form and Stabilize Water-in-Oil Emulsions (the Emulsions of the Invention)

[0259] Unexpectedly, we were able to form 10% water-in-oil emulsions (reverse emulsions) using 1% (w/w) non-micronized curcuma extract particles (FIG. 14A) or 2.5% (w/w) non-micronized rosemary leave extraction cake particles (FIG. 14B)

Example 12. Characterization of the Ability of Particles Prepared from One or More Biological Materials where the Biological Material is Obtained from a Photosynthetic Organism to Form and Stabilize Water-in-Oil Emulsions (the Emulsions of the Invention) after that the Particles have been Washed with Water

[0260] When washing the natural particles with water, we have unexpectedly found that the resulting supernatants exert, for most of them, a significant, although relatively modest, tensio-activity (i.e. the ability to decrease the tension at the interface between stripped sunflower oil and water (FIG. 15)). Hence, to decipher if the stabilizing effect previously seen in Examples 6, 7, and 8 was merely due to the tensio-activity of some surface-active molecules contained in the powders and removable by washing or was more specifically due to a Pickering (mechanical) stabilization mechanism, we recapitulated some physical stability tests on Pickering emulsions stabilized by the washed particles. Interestingly, the washed particles were all able to physically stabilize the resulting oil-in-water emulsions (FIG. 17). We also tested the supernatant resulting from the washing procedure and found that they were not able to stabilize oil-in-water emulsions (FIG. 16), thus clearly showing that the stabilizing effect is conveyed by a true Pickering mechanism and not by a conventional emulsifying effect.

Example 13. Comparison of Sunflower Oil-in-Water Emulsions where the Antioxidant (α-Tocopherol) is Either (i) in the Palmitin Colloidal Particles (the Emulsion of the Invention Prepared as Detailed in the Materials and Methods Section) or (ii) within the Interior of the Oil Droplets

[0261] Two Pickering emulsions prepared as detailed above were prepared. One with a-tocopherol incorporated in the colloidal particles (FIG. 18a) and one with α-tocopherol in the liquid sunflower oil droplets (FIG. 18b).

[0262] Oxidation was accelerated with 200 μM FeSO.sub.4/EDTA at 25° C. The emulsions were then tested for both primary oxidation products such as conjugated dienes and secondary oxidation products such as p-anisidine aldehydes. As shown in FIGS. 18c and 18d, less oxidation occurred in the emulsion when α-tocopherol was incorporated in the colloidal particles compared to the same emulsion where α-tocopherol was solubilized in the interior of the oil droplets.

[0263] The stability of α-tocopherol in each emulsion was then investigated by extracting a-tocopherol from either the colloidal particles or emulsion droplets as described in FIGS. 18a and b.

[0264] HPLC analysis of the α-tocopherol showed that incorporating α-tocopherol into the colloidal particles provided significant protection to the antioxidant (FIG. 18e).

[0265] This unexpected effect brings an additional advantage to the formulation studied here since α-tocopherol in particular, and phenol-bearing compounds in general, are very sensitive to oxidation mediated by lipid oxidation products such as free radicals.

[0266] To further characterize the improvement in antioxidant activity of α-tocopherol when formulated in colloidal particles (FIGS. 18c and d) along with the protection effect of such a formulation on tocopherol itself (FIG. 18e), confocal laser scanning microscopy at different time intervals (0, 6, 24, and 72 hours) was used to image the emulsion labelled with a fluorescent analog of α-tocopherol (25-NBD-cholesterol) located in the colloid particles (FIG. 19A).

[0267] As it can be seen in FIG. 19A, when the fluorescent dye is in the colloidal particles attached at the interface (i.e. Pickering particles), a strong green fluorescence is present around all lipid droplets at 0 min, forming a ring-like pattern. This shows that a significant part of the α-tocopherol fluorescent analog is located at the interface within adsorbed colloidal particles. With time, α-tocopherol is only slowly released from the colloidal particles to the liquid droplets. This can be seen through the decrease of the black-to-green droplet ratio from 0 to 72 hours.

[0268] In contrast, when the fluorescent analog of α-tocopherol is in the liquid oil droplet (FIG. 19B), all droplets are green, demonstrating that the fluorescent analog of α-tocopherol immediately reaches a dynamical equilibrium within all oil droplets.

[0269] This shows that incorporation of the anti-oxidant (i.e. α-tocopherol) within the colloidal particles allows the anti-oxidant (i.e. α-tocopherol) to locate at the interface and provide an improved anti-oxidant effect whilst being protected by the colloidal particles, and that the anti-oxidant (i.e. α-tocopherol) is only slowly released from the colloidal particles, thus maintaining the anti-oxidant effect.

[0270] Finally, polarized light microscopy (FIGS. 19C and 19D) and differential scanning calorimetry (DSC) analyses (FIG. 20) showed that the colloidal particles adsorbed at the oil-water interface, remained physically intact over the timescale of the experiment. This result suggests that the diffusion of α-tocopherol fluorescent analog (hence, by analogy, the diffusion of α-tocopherol) is caused by the solubilization of the colloidal particles in the liquid oil phase over time, which is in line with the high long-term physical stability of those emulsions.

Example 14. Comparison of Sunflower Oil-in-Water Emulsions where the Antioxidant (Carnosic Acid) is Either (i) in the Palmitin Colloidal Particles (the Emulsion of the Invention Prepared as Detailed in the Materials and Methods Section) or (ii) within the Interior of the Oil Droplets

[0271] The exact same experimental design as Example 13 was reproduced except carnosic acid was used to replace α-tocopherol. The data presented in FIG. 21 clearly showed that the same antioxidant effect is obtained with this diterpene phenolic antioxidant, indicating that the colloidal particles of the invention can serve as interfacial reservoirs for many antioxidant molecules to provide an antioxidant-enhancing effect.

Example 15. Comparison of Sunflower Oil-in-Water Emulsions where the Antioxidant (α-Tocopherol) is Either (i) in Palmitin Colloidal Particles not Adsorbed at the Interface or (ii) is within the Interior of the Oil Droplets

[0272] To investigate whether the improvement of the antioxidant activity of α-tocopherol formulated in colloidal particles was specifically due to the interfacial anchorage of these particles, two types of conventional sodium caseinate-stabilized stripped sunflower oil-in-water emulsions comprising an aqueous suspension of colloidal particles were prepared.

[0273] The emulsion and suspension compositions were identical, but the antioxidant was located either in the suspended colloidal particles (FIG. 22a) or in the core of the oil droplets (FIG. 22b). The main difference compared to Example 3 was that the colloidal particles were not adsorbed to the interface and were instead added as a particle suspension in the aqueous phase (i.e. unadsorbed).

[0274] Oxidation was accelerated with 200 μM FeSO.sub.4/EDTA at 25° C. The emulsions were then tested for both primary oxidation products such as conjugated dienes and secondary oxidation products such as p-anisidine aldehydes. As shown in FIGS. 22c and 22d, the oxidative stability of both emulsions was quite similar, and both emulsions contained a higher concentration of oxidation products than the emulsion of Example 3, an emulsion of the invention where the anti-oxidant is contained within the colloidal particles attached to the oil/water interface.

[0275] Extraction of α-tocopherol from the unadsorbed colloidal particles or emulsion droplets, followed by HPLC showed the protective effect conferred by the colloidal particles in Example 13 (FIG. 18e) was dramatically reduced when the particles were not absorbed at the droplet surface (FIG. 22e).

[0276] Again, this shows that the emulsion of the invention (Examples 13 and 14) provides a much better protection to the antioxidant compounds than the two described emulsions of Examples 15. The attachment of the antioxidant-loaded particles to the interface (i.e. true Pickering particles) is thus required to have a beneficial effect in terms of antioxidant activity.

[0277] Finally, laser scanning microscopy at different time intervals (0, 6, 24, and 72 hours) was used to image the emulsion labelled with a fluorescent analog of α-tocopherol (25-NBD-cholesterol) located in the un-adsorbed colloid particles (FIG. 23A) or directly in the oil droplets (FIG. 23B).

[0278] As it can be seen, when the fluorescent dye is in the suspended particles, a green fluorescence is homogenously distributed in the external aqueous phase at 0 min. At that incubation time, no green droplets can be observed. This shows that at 0 hour, no colloidal particles are adsorbed at the oil/water interface. Instead they are suspended in the aqueous phase as above-mentioned where they are quite inefficient to counteract lipid oxidation. With time, a depletion of the green background is paralleled by an increase of the green droplet, clearly showing that the fluorescent analog of α-tocopherol progressively diffuse from the aqueous phase to the oil droplet interior.

[0279] At 72 hours, the black-to-green droplet ratio is similar (FIG. 22A), if not identical, as the one depicted in FIG. 23B where the dye was initially located in the oil droplet core.

Example 16. Comparison of Pickering Sunflower Oil-in-Water Emulsions where the Antioxidant (α-Tocopherol) is Either (i) in Palmitin Colloidal Particles not Adsorbed at the Interface and Refrained to Diffuse in the Oil Interior by a “Pickering Barrier” or (ii) is within the Interior of the Oil Droplets

[0280] To decipher the respective contribution of each population of colloidal particles (adsorbed vs. non-adsorbed at the interface) in the enhancing effect on α-tocopherol antioxidant activity seen in Example 13 (the emulsion of the invention), a Pickering emulsion stabilized by α-tocopherol-free colloidal particles attached to the interface was prepared, to which a-tocopherol-loaded colloidal particles were added post homogenization, resulting in the a-tocopherol-loaded colloidal particles not attaching to the interface (FIG. 24A).

[0281] This emulsion was compared to the same Pickering oil-in-water emulsion except the a-tocopherol was located in the core of the oil droplets (FIG. 24B).

[0282] This comparison allowed the assessment of the role of antioxidant-loaded CLPs in the continuous phase, while keeping the interfacial structure similar to that of the emulsion of the invention (Example 13).

[0283] Lipid oxidation proceeded significantly faster in Pickering emulsions containing a-tocopherol exclusively in the colloidal lipid particles (FIGS. 24C and 24D). Thus, we can conclude that the antioxidant-loaded particles suspended in the aqueous phase of the emulsion of the invention (Example 13) have no contribution to the antioxidant activity-improving effect. Indeed, here it is clearly shown that this precise antioxidant-loaded particle subpopulation is less effective at inhibiting lipid oxidation than α-tocopherol directly formulated in the oil phase.

[0284] As in previous Examples, emulsions with similar construction principle were also prepared with 25-NBD cholesterol. The diffusion of the fluorescent probe from the colloidal particles present in the aqueous phase to the emulsion droplet core during incubation was much slower compared to the protein-stabilized emulsion (FIG. 25A), which confirms the beneficial effect that the colloidal particles provide to the emulsions of the invention as seen in Example 13.

Example 17. Comparison of Pickering Sunflower Oil-in-Water Emulsions with Different Concentrations of an Antioxidant (α-Tocopherol) which is Located Either (i) in the Tripalmitin Colloidal Particles (the Emulsion of the Invention Prepared as Detailed in the Material and Methods Section) or (ii) within the Interior of the Oil Droplets

[0285] To investigate to what extent our hierarchical emulsion design presented in Example 13 boosted the antioxidant efficiency of an antioxidant as compared to a control emulsion where the antioxidant is located within the interior of the oil droplets, an emulsion of the invention was produced with a reduced α-tocopherol content from 90 ppm to 45, then 22.5 ppm (2 to 4 times lower).

[0286] Interestingly, we found that, when formulated in the emulsion of the invention, tocopherol can be drastically reduced (at least 2 to 3 times) and still provide a protection against lipid oxidation which is superior or equal to that obtained with 2 to 3 times higher concentrations of α-tocopherol in the control emulsion where the antioxidant is formulated in the interior of the lipid droplets (FIG. 26).

Example 18. Physical and Morphological Characterization of Tripalmitin Colloidal Particles with or without Antioxidant (α-Tocopherol)

[0287] In this example, we characterized the particles containing or not α-tocopherol using particle size distribution (A), differential scanning calorimetry (A) and TEM (A) (FIGS. 27A, B and C).

Example 19. Physical and Morphological Characterization of Pickering Sunflower Oil-in-Water Emulsions with Antioxidant (α-Tocopherol) Either in Palmitin Colloidal Particles (the Emulsion of the Invention) or in the Core of the Oil Droplets

[0288] In this example, we characterized Pickering oil-in-water emulsions by measuring the droplet size distribution (A), their thermal properties of melting and crystallization (B), as well as their morphology (C) (FIGS. 28A, B and C).

Example 20. Comparison of the Activity of α-Tocopherol-Loaded Colloidal Particles in the Sunflower Oil-in-Water Emulsions of Example 13 with an Emulsion Prepared Using a Non-Oxidizable Oil

[0289] To investigate whether α-tocopherol-loaded colloidal particles adsorbed at the droplet surface of Pickering emulsion prevents oxidation in the emulsion through a specific antioxidant action and not any other mechanism, the experimental set up of Example 3 was reproduced, except that the oil used in the emulsion consisted of medium chain triglycerides (MCTs) instead of stripped sunflower oil.

[0290] Unlike sunflower oil, MCTs are non-oxidizable and it can be seen in FIG. 29 that there was no significant consumption of α-tocopherol observed in either emulsion, even in the presence of ferrous iron (200 μM FeSO.sub.4/EDTA at 25° C.). This suggests that α-tocopherol is specifically consumed by lipid oxidation products and not directly by cation metals. Consequently, α-tocopherol-loaded colloidal particles exert a protecting effect towards Pickering emulsions through a true antioxidative action.

Example 21. Calorimetrical Characterization of a Conventional Sodium Caseinate-Stabilized Sunflower Oil-in-Water Emulsion Containing Added Palmitin Colloidal Particles in the Aqueous Phase (Solid Line), and a Colloidal Lipid Particles Dispersion (Dashed Line)

[0291] In this example, we characterized a conventional sodium caseinate-stabilized emulsion using differential scanning calorimetry (FIG. 30).

Example 22. Comparison of Sunflower Oil-in-Water Emulsions where the Antioxidant (α-Tocopherol) is Either (i) in the Palmitin (80%) Colloidal Particles Containing 20% Tricaprylin (the Emulsion of the Invention Prepared as Detailed in the Materials and Methods Section) or (ii) within the Interior of the Oil Droplets

[0292] The exact same experimental design as Example 13 has been reproduced here but with colloidal particles containing 20% tricaprylin/80% tripalmitin instead of 100% tripalmitin (FIGS. 31A and B). The data presented in FIGS. 31C, D, and E clearly showed that the same type of advantage is obtainable with these colloidal particles, indicating that the antioxidant-enhancing effect is robust toward variation of the colloidal particle composition.

Example 23. Comparison of Flaxseed Oil-in-Water Emulsions where the Antioxidant (α-Tocopherol) is Either (i) in the Palmitin Colloidal Particles (the Emulsion of the Invention Prepared as Detailed in the Materials and Methods Section) or (ii) within the Interior of the Oil Droplets

[0293] The exact same experimental design as Example 13 has been reproduced here but with flaxseed oil instead of sunflower oil (FIGS. 33A and B). The data presented in FIG. 33C clearly showed that the same type of advantage is obtainable with a different oil than sunflower oil, indicating that the antioxidant-enhancing effect is robust toward variation of the oil composition.