Antioxidant loaded emulsions entrapped in liposomes produced using a supercritical assisted technique

J. of Supercritical Fluids 154 (2019) 104626

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Antioxidant loaded emulsions entrapped in liposomes produced using a supercritical assisted technique P. Trucillo a , R. Campardelli a,b,∗ , E. Reverchon a a b

Department of Industrial Engineering, University of Salerno, Via Giovanni Paolo II, 132, 84084, Fisciano, Italy Department of Civil, Chemical and Environmental Engineering (DICCA), University of Genoa, Via Opera Pia 15, 16145, Genova, Italy

h i g h l i g h t s

g r a p h i c a l

a b s t r a c t

• A novel approach to include lipophilic molecules in liposomes using emulsions. • Production of stable liposomes loaded with antioxidant loaded emulsions. • Encapsulation Efficiencies of antioxidants up to 99%. • High preservation of antioxidant power using emulsion loaded liposomes.

a r t i c l e

i n f o

Article history: Received 4 July 2019 Received in revised form 14 August 2019 Accepted 7 September 2019 Available online 11 September 2019 Keywords: Liposomes Antioxidants Molecule preservation Emulsions Supercritical fluids

a b s t r a c t Two different strategies for the encapsulation of hydrophobic bioactives within nanoliposomes were attempted. Three model lipophilic antioxidants (farnesol, limonene, linalool) were entrapped in liposomes using emulsions: drug was dissolved in the oil phase of an oil in water emulsion; then, the aqueous emulsion was entrapped in liposomes inner core using SuperLip (Supercritical assisted Liposome formation). Vesicles loaded with antioxidants in the lipidic layer, showed mean sizes from 116 ± 32 nm to 230 ± 62 nm, with an Encapsulation Efficiency (EE) up to 74% for farnesol, 87% for limonene and 54% for linalool. The use of O/W emulsion, resulted in larger liposomes (among 397 ± 103 nm and 605 ± 175 nm), but also in an EE up to 99%. O/W emulsion method successfully preserved the antioxidant activity of the entrapped molecules. The reduction of the antioxidant power was about 22% for lipidic layer entrapment; whereas, only 2% using O/W method. © 2019 Elsevier B.V. All rights reserved.

1. Introduction Abbreviation: PC, l-alpha-phosphatidylcholine; PSD, particle size distributions; DLR, drug to lipid ratio; EE, encapsulation efficiency; Zpot, zeta potential; PDI, polydispersity index; O/W, oil in water; FE-SEM, field emission-scanning electron microscope; Lin, linalool; Far, farnesol; Lim, limonene; DPPH, 2,2-diphenyl-1picrylhydrazyl; DSD, droplet size distribution. ∗ Corresponding author at: Department of Industrial Engineering, University of Salerno, Via Giovanni Paolo II, 132, 84084, Fisciano, Italy. E-mail address: [email protected] (R. Campardelli). https://doi.org/10.1016/j.supflu.2019.104626 0896-8446/© 2019 Elsevier B.V. All rights reserved.

Liposomes are spherical colloidal systems [1] formed by one or more double layers of phospholipids surrounding an inner aqueous core [2]. These lipidic vesicles attracted the scientific and industrial attention [3], for their biocompatibility with human cell barrier [4]. Indeed, liposomes can reach the target cell and fuse with its membrane, releasing the drug content directly into the cytoplasm, avoiding molecule leakage and toxic side effects [5]. Liposome have

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been proposed in several industrial fields such as food [6], cosmetic [7], pharmaceutic [8] and textile [9]. They are versatile drug delivery systems [10], since they can be employed to transport hydrophilic molecules in the inner core [11], or lipophilic molecules in the double lipidic layer [12]. The encapsulation of bioactive molecules into liposomes, guarantees several advantages such as the enhancement of drug bioavailability [13], protection from degradation and therapeutic efficacy [14]. Conventional liposomes fabrication processes are mainly based on the dissolution of lipids into an organic solvent; then, a thin layer of phospholipids is obtained and subsequently hydrated using an aqueous phase containing the drug [15]. However, this method is characterized by several drawbacks, such as low encapsulation efficiency (20–30 %), large particles size distribution and low process replicability due to batch configuration [16]. A process that causes high solvent residue in the final product is ethanol injection [17]. To avoid these drawbacks, time consuming and high expensive post-processing steps have been proposed, such as extrusion [18] or sonication [19]. Several supercritical assisted techniques have been developed in the last decades to provide alternatives to the traditional processes for liposomes production, such as Supercritical Fluid Extraction [20], Supercritical Anti-Solvent [21], Supercritical Reverse Phase Evaporation [22] and Depressurization of an Expanded Solution into an Aqueous Medium [23]. However, all these processes only partially resolve the problems in liposomes production, since not a good control of particle size distribution and low encapsulation efficiencies are still a challenge [24]. Therefore, a supercritical assisted process named Supercritical assisted Liposome formation (SuperLip) was recently developed to obtain a large scale production of lipidic vesicles [25]. Nanometric liposomes were produced with particle size distributions narrower than conventional processes and encapsulation efficiencies high up to 99% [26]. The idea at the basis of the SuperLip process consisted in the inversion of the traditional production steps of liposomes: first, droplets of water were atomized in a Formation Vessel (FV of Fig. 1), obtaining a water in carbon dioxide emulsion; then, lipids were rapidly added to the system.

It was already demonstrated that, in SuperLip, water droplets can be considered as discrete volumes in which the dissolved or suspended molecules are confined during the encapsulation process [27]. This hypothesis was used to entrap water-soluble compounds into the aqueous inner volume of liposomes. Lipophilic drugs were incorporated into the phospholipid bilayer membrane [28]. As reported in this study, an amphiphilic antioxidant compounds such as eugenol was entrapped in both liposomes compartment. In particular, the encapsulation in the most exposed region of the vesicles caused the degradation of the entrapped molecule; whereas, the antioxidant power of the entrapped compound was better preserved when it was included into the inner aqueous core [29]. However, several antioxidant molecules are lipophilic and could be encapsulated only in the double lipidic layer of vesicles. Several attempts at antioxidant encapsulation into liposomes are also reported in literature [30–33], using the strategy of the double layer entrapment, especially in the food and dietary supplement production field [34–37]. A possible different strategy to entrap lipophilic compounds into the lipidic inner core is the incorporation of the compound in a stable Oil in Water (O/W) emulsion that, in a second step, is entrapped into liposomes aqueous core. At the best of the author’s knowledge, the production of emulsions for lipidic molecules inclusion in the aqueous core of liposomal vesicles has never been tested before. For this reasons, in this work, this approach has been proposed and tested. Lipophilic drug has been dissolved into an oil phase of an Oil in Water emulsion (O/W); then, the emulsion has been entrapped in the inner core of liposomes. Lipophilic antioxidants such as limonene (monoterpene not oxygenated), linalool (monoterpene oxygenated) and farnesol (sesquiterpene oxygenated) were selected as model molecules to verify this process. Moreover, the simultaneous entrapment of 2 of these molecules will be also attempted, with the aim of improving the performance of these additives, such as activity, antioxidant power preservation and encapsulation efficiency. The process will be compared with the double layer inclusion method. The preservation of the antioxidant power, liposome size distribution, vesicles morphology and encapsulation efficiency of loaded molecules will be studied. 2. Materials, apparatus and methods 2.1. Chemicals and reagents L-␣-Phosphatidylcholine from egg yolk (PC, 99% pure, powder, CAS 8002-43-5), limonene (97% pure, liquid, MW 136 g/mol, CAS 5989-27-5), linalool (97% pure, liquid, MW 154 g/mol, CAS 78-706), farnesol (95% pure, liquid, MW 222 g/mol, CAS 4602-84-0) were obtained from Sigma Aldrich, Milan, Italy. Ethanol (>99% pure, CAS 64-17-5) was used to dissolve phospholipids (highly soluble at room temperature and atmospheric pressure); it was purchased by Sigma Aldrich, Milan, Italy. The surfactant Tween 80 (CAS 9005-65-6) and the oil Isopropyl myristate (>90% pure, CAS 110-27-0) were obtained by Sigma Aldrich, Milan, Italy, and were used for the emulsions formulation. Carbon dioxide (>99.4% pure) was provided by Morlando Group, Napoli, Italy. Distilled water was self-produced using a lab-scale distillator and used in all liposomes formulations. The reagent 2diphenyl-1-picrylhydrazyl (DPPH, CAS 1898-66-4) was purchased from Sigma Aldrich, Milan, Italy. All the chemicals and reagents have been used as received. 2.2. Apparatus

Fig. 1. SuperLip layout sketch.

Supercritical assisted Liposome formation (SuperLip), is described in the sketch of Fig. 1. Pure carbon dioxide (flow rate

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Fig. 2. Farnesol (a), Limonene (b) and Linalool (c) loaded liposomes at theoretical loadings of 10%, 20% and 30% w/w on lipid mass basis.

of 6.5 g/min, Ecoflow pump, mod. LDC-M-2, Lewa, Germany) is fed together with an ethanolic solution (flow rate of 3.5 mL/min, corresponding to 2.76 g/min, high pressure precision pump, Model 305, Gilson, France) to a mixing element, called saturator (S1 in Fig. 1). Ethanol is used to dissolve phospholipids and lipophilic antioxidants (in case of encapsulation into the lipidic double layer of liposomes). The flux of ethanol and carbon dioxide creates an expanded liquid in the saturator (internal volume of 0.3 dm3 ), and is loaded with 5 mm2 Berl saddles. The working temperature of the saturator is set at 40 ◦ C; whereas, the pressure is set at 100 bar. Ethanol and CO2 feed rates are selected to obtain a Gas to Liquid Ratio of 2.4 on mass basis, as already optimized in a SuperLip previous work [38]. The expanded liquid, i.e. a liquid mixture that contains a large amount of the dissolved gas, is then delivered to a high pressure formation vessel (FV in Fig. 2, internal volume of 0.5 dm3 ), working at the temperature of 40 ◦ C and pressure of 100 bar). An aqueous solution (the third feeding line in Fig. 1), or an O/W emulsion in which the antioxidant compound has been previously stabilized, is delivered at a flow rate of 10 mL/min to the formation vessel (FV) using a high pressure precision pump, Model 305, Gilson, France. The aqueous solution (or the emulsion) is sprayed into the formation vessel (FV) using an 80 ␮m internal diameter nozzle. The liposomes are produced in aqueous suspension, that is temporary stored in a unit set at the bottom of the formation vessel, called Accumulation Unit (A, in Fig. 1). The organic solvent used for the expanded liquid production is recovered by the expansion in a separator (S2 in Fig. 1), working at reduced pressure (10 bar). To avoid Joule Thomson effect, the piping line that links the FV and the S2 is heated at 30 ◦ C. Carbon dioxide flow rate is measured using a rotameter (mod. N.5–2500, Serval 115022, ASA, Italy); whereas, liposomes are collected from the bottom of the formation vessel in an aqueous bulk (A), at regular interval times, using an on/off valve. Operative parameters such as Pressure, Temperature, Gas to Liquid Ratio of the Expanded Liquid and Water Flow rate were selected from previous optimization studies [28,38–42].

2.3. Antioxidant encapsulation tests procedure 3 different antioxidants theoretical loadings (10%, 20% and 30%) on lipid basis were tested for liposomes entrapment. Lipid and antioxidant were both dissolved in the ethanol phase using the method of inclusion in the double lipid layer. When the emulsion method was tested, oil in water O/W emulsion in the ratio oil/water 10/90 were prepared following this procedure: the oil phase (10 g) was prepared dissolving the selected antioxidant in isopropyl myristate. The water phase (90 g) was obtained dissolving surfactant Tween 80 at 0.2% w/w in distilled water. Then, the emulsions were obtained with an emulsifier (mod. L4RT, Sil-

verstone, USA), working at 7000 rpm for 6 min. Emulsions were directly processed using SuperLip, to avoid antioxidant loss due to high volatility of the chosen compounds. 2.4. Methods 2.4.1. Liposomes and emulsions mean size Mean size and polydispersity index of O/W emulsions were measured using Dynamic Light Scattering (Mastersizer S, Malvern Instruments Ltd., Worcherstershire, UK). Analyses were performed 30 min after emulsion preparation. However, a volume of the O/W emulsion was separated and analyzed using DLS also after 24 h, to confirm emulsion stability. After the SuperLip process, the produced lipidic vesicles were analyzed using Dynamic Light Scattering (DLS, Mod. Zetasizer Nano S, Worcestershire, UK), to measure mean diameter (MD) standard deviation (SD) and to compare Particle Size Distributions (PSD). Measurements were performed in triplicates. Liposomes prepared using SuperLip process, as an aqueous suspension, were also previously tested for stability with time [40]. 2.4.2. Optical and Scanning electron microscope observations The emulsions were also observed using an optical microscope (OM, mod. BX 50 Olympus, Tokyo, Japan), equipped with a phase contrast condenser. The 3D-morphology of lipidic vesicles was observed using a Field Emission-Scanning Electron Microscope (FE-SEM, model LEO 1525, Carl Zeiss SMT AG, Oberkochen, Germany). The samples were collected in the liquid suspension after SuperLip process and prepared for the observation: a drop of liposome suspension was spread on an aluminum stub and left drying for 48 h. Then, the samples were covered with gold using a sputter coated at a thickness of 250 Å (model B7341, Agar Scientific, Stansted, United Kingdom). 2.4.3. Encapsulation efficiency To calculate the Encapsulation Efficiency (EE), liposomes liquid suspensions were centrifuged at 6500 rpm per 40 min and the supernatant was separated from pellet. Then, two methods were used for the determination of antioxidant EE: in the first case, the absorbance of antioxidant dissolved in the supernatant was measured, using a method reported in literature [43,44]. For this measurement, a Micro-volume UV–vis spectrophotometer (BioSpec-nano, Shimadzu Scientific Instruments, Columbia, USA) has been used. The EE was calculated using the following equation:

EE [%] =

ppmtheoretical − ppmsupernatant ppmtheoretical

*100

in which ppmtheoretical is the amount of drug dissolved in the oil phase (ethanol or isopropyl myristate) at the beginning of the process. Instead, ppmsupernatant represents the not entrapped

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antioxidant; i.e., the free drug in the external aqueous bulk of liposomes suspension. Spectra of free linalool, limonene and farnesol were acquired from the literature [45–50]; then, the maximum absorbance peaks were detected at 220 nm for linalool, 270 nm for limonene and 260 nm for farnesol. For this reason, the simultaneous encapsulation of 2 compounds was performed only for linalool + limonene and for linalool + farnesol systems, to avoid the overlap of detected peaks. Entrapment efficiencies measurements have been performed in triplicates and the results are the mean of the calculated efficiencies. As a confirmation of the data obtained for the supernatant, the encapsulation efficiency was also measured from the pellet; i.e., the centrifuged lipidic vesicles. After centrifugation and pellet separation, liposomes were dissolved in 2 mL ethanol, disrupting the lipidic barriers of liposomes. Then, the absorbance of the sample was measured, using as a blank the solution ethanol + isopropyl myristate + lipids at the same concentration of the experiments. The EE was then calculated using the following equation EE [%] =

ppmpellet ppmtheoretical

*100

and the results overlapped the ones obtained with the supernatant method. EE determination was performed in triplicates for each experiments. Results reported in the Results section are a mean of the value obtained from both methods. 2.4.4. Antioxidant activity Antioxidant activity of the compounds entrapped in lipidic vesicles was measured as the radical-scavenging ability by means of 2,2-diphenyl-1-picrylhydrazyl (DPPH•). Following the preparation protocol, 0.0197 g of DPPH were dissolved in 500 mL ethanol to obtain a 10−4 M DPPH ethanol solution [51]. According to PubChem and Sigma Aldrich databases, the solubility of linalool in water is 1.59 mg/mL, limonene solubility in water is 13.8 mg/L and is 1.7 mg/L for farnesol. To obtain a calibration line, 4 diluted suspensions (50, 100, 200 and 500 ppm) were obtained and 1 mL of each calibration sample was mixed with 3 mL of DPPH in ethanol solution. The mixtures were stored in glass for 30 min in the dark. Then, the UV–vis spectrophotometer was used at a wavelength of 517 nm. In the case of molecules entrapped directly into the lipidic layer of liposomes, the suspensions were centrifuged at 6500 rpm for 40 min at −4 ◦ C and the lipidic pellet was separated from the supernatant. Then, the pellet (i.e., the liposomes containing the antioxidant) was dissolved in ethanol and DPPH method was applied to disrupt vesicles membranes and let the molecules to diffuse into the external bulk. The DPPH inhibition percentage was calculated according to the following equation:



Inhibition [%] = 100* 1 −

Aa Ab



in which Aa is the absorbance (obtained at 517 nm) of the sample treated with DPPH in ethanol solution, and Ab is the control absorbance (at 517 nm) of DPPH in ethanol. The inhibition percentage was then compared to the one of pure unprocessed samples percentage, to calculate the decrease of DPPH inhibition capacity between unprocessed and processed samples. Measurements have been performed in triplicates and the results are the means of the calculated inhibition percentages. 3. Results and discussion In previous works performed with SuperLip process, it has been demonstrated that using this process it is possible to atomize a hydrophilic solution and efficiently encapsulate it into liposomes. In the following section, experimental results obtained in this

research are presented and discussed with the objective of demonstrating that also an emulsion can be entrapped into liposome core using this process. For this reason, lipophilic antioxidants will be stabilized in O/W emulsions that will be then processed using SuperLip. Lipidic layer entrapment experiments were also performed for comparison purposes. A first set of experiments was performed to entrap farnesol, linalool or limonene in the double lipidic layer of liposomes; i.e., in the external lipophilic compartment of the vesicles. The first lipid layer encapsulation experiments were performed varying antioxidant theoretical loading in the feeding, on lipidic mass basis (10%, 20% and 30% w/w), as reported in Table 1. Particle Size Distributions (PSDs) of farnesol, limonene and linalool loaded liposomes are shown in Fig. 2a–c. Fig. 3 shows that liposomes with a good control on size distribution were obtained; the distribution curves practically overlap by increasing the antioxidant loading. Nanometric liposomes were obtained. As reported in Table 1, mean diameter of farnesol loaded liposomes ranged between 126 ± 35 nm for 10% theoretical loading and 146 ± 44 nm for 30% loaded liposomes (see also Fig. 2a). The PolyDispersity Indexes (PDIs) ranged between 0.27 and 0.34, indicating that narrow size distributions have been obtained for all the loadings tested. However, the encapsulation efficiency (EE) was low, around 22%, for the experiment performed at the lowest farnesol concentration; whereas, an improvement of antioxidant EE was observed increasing theoretical loading. Indeed, an EE of 65% was obtained in correspondence of the experiments performed at 20% w/w of theoretical loading and an EE of 74% for 30% w/w farnesol theoretical loading. The same encapsulation experiments were performed using limonene (see the results in Table 1). Limonene loaded liposomes mean diameters are included between 116 ± 42 nm and 159 ± 32 nm, also in this case the effect of the increase of the antioxidant loading on liposome mean diameters was negligible. A good control of liposomes size distribution was obtained also in this case (see Fig. 2b), with PDIs among 0.28 and 0.35. Looking at data showed in Table 1, high EEs were obtained; it is possible to observe an increasing trend, also in this case from 67% of EE (10% w/w) to 87% (30% w/w), when increasing limonene theoretical loading. Linalool was also entrapped inside liposomes at the same process conditions (see Table 1 and Fig. 2c) reported in the Apparatus section. Looking at data reported in Table 1, it is possible to observe that liposome mean diameter of 197 ± 62 nm was obtained in the case of 10% w/w linalool theoretical loading; whereas, liposomes with 230 ± 62 nm mean diameter were observed in correspondence of 30% loading, with a PDI among 0.27 and 0.32. Encapsulation Efficiencies were lower with respect to the other compounds tested (from 35% to 54%). However, an increase of Encapsulation Efficiency can be observed with the increase linalool theoretical loading. Regarding the encapsulation tests, these results can be probably due to the low molecular affinity among linalool and phospholipids, maybe determined by Van Der Walls repulsion forces between lipids and antioxidant molecules [52]. Simultaneous encapsulation of the antioxidants was attempted to verify the feasibility of an antioxidant mixture entrapment in liposomes double layer. Farnesol and linalool together were first entrapped into liposomes with a theoretical loading of 30% w/w (farnesol to lipid ratio) and 30% w/w (linalool to lipid ratio). From Table 1, it can be observed that the simultaneous encapsulation experiment produced liposomes with mean diameter of 181 ± 51 nm and the PDI was equal to 0.28. Encapsulation efficiency was 61% for linalool and 70% for farnesol, respectively. Linalool entrapment efficiency was almost similar than in single entrapment, varying from 54% to 61%. In the case of farnesol, the same

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Table 1 Mean diameter, polydispersity index, and encapsulation efficiency of liposomes loaded with farnesol, limonene or linalool in the double lipidic layer. Compound

Farnesol

Limonene

Linalool Lin + Far Lim + Lin

Theoretical loading [%, w/w]

Mean Diameter [nm ± SD]

PDI

EE [%]

10 20 30 10 20 30 10 20 30 30 ; 30 30 ; 30

126 ± 35 132 ± 45 146 ± 44 121 ± 42 116 ± 32 159 ± 44 197 ± 62 213 ± 68 230 ± 62 181 ± 51 121 ± 44

0.30 0.34 0.27 0.35 0.28 0.28 0.31 0.32 0.27 0.28 0.36

22 65 74 67 87 87 35 36 54 Lin 61 Lim 90

Far 70 Lin 61

Fig. 3. FE-SEM images of double lipidic layer loaded liposomes containing (a) farnesol and (b) limonene.

entrapment efficiency of about 70% was obtained, compared to the single entrapment experiment (74%). A second mixture of antioxidants limonene-linalool was entrapped in the double lipidic layer of liposomes, maintaining constant the operative parameters of the previous experiment. Liposomes with mean diameter of 121 ± 44 nm and PDI of 0.36 were obtained. Also in this case, the simultaneous entrapment efficiency of the two compounds resulted practically unchanged. Indeed, in the simultaneous entrapment experiments, limonene was entrapped at 90% and linalool at 61%; whereas, they were entrapped with an EE of 87% and 54% respectively in the single entrapment experiments. Fig. 3a-b is related to the FE-SEM observations of liposomes obtained with the lipidic layer entrapment method. As reported in Fig. 3a-b, liposomes are characterized by nanometric and sub-micrometric dimensions, showing spherical and smooth surface. Distribution of the vesicles is homogenous. The proposed images of farnesol (Fig. 3a) and limonene (Fig. 3b) loaded samples do not show significant differences varying the entrapped compound. In the second part of this work, the chosen antioxidant compounds were entrapped into O/W emulsions, adopting the strategy described in the Methods section. The basic principle of this procedure was to first stabilize the antioxidant into O/W emulsion; then, this emulsion was atomized in the SuperLip formation vessel to obtain liposome with emulsion included in the inner core. Emulsions stability was first studied; experiments were performed for the entrapment of limonene and linalool into O/W emulsions. The antioxidants were dissolved in isopropyl myristate (oil phase) and, then, emulsified with a water surfactant solution. Fixed amount of limonene and linalool were dissolved in the oil phase, in order to respectively obtain 10%, 20% and 30% feeding loading ratio over phospholipid mass. Emulsions obtained were stable during time with non-coalescing droplets. An optical microscope image of the obtained emulsion is reported in Fig. 4.

Fig. 4. Optical image of limonene loaded emulsions.

Table 2 Mean diameters and polydispersity indexes of droplets obtained by oil in water emulsions loaded with limonene and linalool. Compound

Limonene

Linalool

Theoretical loading [%, w/w]

Mean Diameter [␮m ± SD]

PDI

10 % 20 % 30 % 10 % 20 % 30 %

4.1 ± 1.4 3.9 ± 1.3 3.7 ± 0.8 3.5 ± 1.6 3.4 ± 1.3 2.5 ± 0.8

0.36 0.33 0.21 0.46 0.38 0.33

Emulsions were characterized by micrometric droplet dimensions, as reported in Table 2. Treatment of the emulsion in a supercritical system was investigated. For this reason, the previously prepared emulsion was atomized in the high pressure formation vessel (see the process

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Fig. 5. Particle Size Distributions of emulsion before and after the contact with the supercritical mixture in the SuperLip formation vessel.

sketch of Fig. 1), filled with an ethanol/CO2 mixture; i.e., without the addition of phospholipids (PC). In this manner, liposome could not be formed. The emulsions prepared in this manner were collected at the bottom of the SuperLip plant and their Droplet Size Distribution (DSD) was analyzed. The comparison of Droplet Size Distribution of the emulsion before and after the contact with the supercritical mixture is reported in Fig. 5. In this figure, a re-arrangement of the emulsion after SuperLip process is evident: sub-micrometric and nanometric level were obtained, with narrower particle size distributions. This result is a consequence of fluid dynamics solicitation on the emulsion in the SuperLip process. The mechanisms involved in droplets re-arrangement could depend on several parameters, such as atomization in a high pressure environment [53], turbulent shear forces and viscosity of the medium [54]. Emulsion droplets could undergo mitosis-like disruption [55]. Indeed, high pressure jet technique is commonly used to obtain emulsion droplets at nanometric level; the higher is the energy density of the atomization [56], due to nozzle micrometric diameters and high pressure, the smaller are the droplets produced [57]. For these reasons, it was not necessary to provide post-processing steps for obtaining emulsions at nanometric level, since the final dimensions were reached thanks to the jet break up due to the high pressure set in the Formation Vessel (FV of Fig. 1). Once confirmed that the emulsion was not destabilized during atomization in the formation vessel and smaller droplets were even produced, a complete SuperLip experiment was performed; i.e., involving the lipids in the system. Emulsions were

atomized in the formation vessel containing the ternary mixture CO2 /ethanol/Lipids to obtain the formation of liposomes containing the atomized emulsion droplets. The experiment were performed successfully and liposome stable suspensions loaded with limonene and linalool emulsions were recovered at the bottom of the plant (see results reported in Table 3). Analyzing the results obtained in these experiments, a first observation is that liposome mean diameter was considerably smaller than emulsion droplets, confirming the result obtained in the preliminary experiment of emulsion processing. Since in the formation vessel also phospholipids are present, sub-micrometric liposomes were obtained containing the atomized emulsion inside the aqueous core of the vesicles (see Fig. 6a-b). Liposomes loaded with limonene emulsions were 397 ± 103 nm large (see Table 3). Encapsulation Efficiency (EE) of limonene entrapped via emulsion was 93%, that is highly improved with respect to limonene EE obtained using the double layer entrapment strategy (67%). Considering the encouraging results, 20% and 30% limonene loading were attempted via emulsion entrapment. Stable emulsions and, then, stable liposome suspensions were produced in all cases (see Table 3). Moreover, increasing limonene loading, an increase of liposomes mean diameter was observed as also evident from PSD comparison reported in Fig. 6a. Limonene EE was larger than 90% in all these cases. These results confirmed the possibility to include emulsions into liposome inner core. Another set of experiments was performed dissolving linalool inside O/W emulsion (Fig. 6b). The same theoretical loading was obtained with respect to lipid (PC) mass (10, 20 and 30%). Linalool loaded samples showed an increasing trend from 424 ± 165 nm to 605 ± 175 nm (see Fig. 6b). Linalool loaded vesicles showed an encapsulation efficiency of 96%, 97% and 99% for 10%, 20% and 30% antioxidant theoretical loading on lipid mass basis; whereas, it was 35%, 36% and 54% for double lipidic entrapment. Simultaneous entrapment of limonene and linalool resulted in an EE of 99% for both compounds; whereas, it was 90% (limonene) and 61% (linalool) in the case of lipidic compartment encapsulation. Even if the EEs were higher than conventional methods when entrapped in the lipidic layer, they were larger when encapsulated in the inner core, using the novel method of emulsion entrapment. As an example of the obtained liposomes loaded with emulsions, Fig. 7 reports the FE-SEM observation of limonene emulsions loaded liposomes. As reported in this figure, liposomes have sub-micrometric mean size, a smooth and spherical surface and vesicles distribution is still homogenous. The larger standard deviations observed in liposomes loaded with 10% w/w limonene (on mass basis) are probably due to the interaction between the antioxidant molecule and lipids, that in case of low drug concentration resulted in a wider lipid aggregation phenomena. As a general comment, the aggre-

Fig. 6. Limonene (a) and Linalool (b) loaded emulsions entrapped in the inner core of liposomes.

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Table 3 Mean diameter, polydispersity index, and encapsulation efficiency of liposomes loaded with limonene and linalool O/W emulsions in the inner core. Compound

Limonene

Linalool Lim + Lin

Theoretical loading [%, w/w]

Mean Diameter [nm ± SD]

PDI

EE [%]

10 % 20 % 30 % 10 % 20 % 30 % 30 % + 30 %

397 ± 103 492 ± 148 655 ± 218 424 ± 165 521 ± 167 605 ± 175 489 ± 117

0.33 0.30 0.26 0.39 0.32 0.29 0.24

93 91 92 96 97 99 99 Lim

99 Lin

Table 4 Antioxidant activity test in different compartment of the produced liposomes. Compound

Linalool

Limonene

Theoretical loading [%, w/w] 10 20 30 10 20 30 10 20 30 10 20 30

Fig. 7. FE-SEM image of inner core loaded liposomes containing limonene O/W emulsion.

gation of liposomes could be also due to the preparation of stubs followed by metallization pre-processing for microscopic observations (dried samples). Instead, the aggregation phenomena are not evident in the optical image (Fig. 4), since the observation occurred in a liquid bulk and the sample was opportunely diluted before observation. 3.1. Antioxidant activity measurement Encapsulation efficiency was higher in the case of Oil-in-Water entrapment into liposomes than in double lipidic layer encapsulation. A compound entrapped in the external lipidic layer is easily exposed to degrading agents such as the high pressures of the system or the temperature. To verify this hypothesis, the reduction of the inhibition power was measured in order to establish if the antioxidant activity is better preserved after the entrapment in the inner core than in lipidic layer. Looking at Table 4, it is possible to observe that linalool entrapped in the double lipidic layer showed an inhibition reduc-

Compartment Lipidic layer

Aqueous core

Lipidic layer

Aqueous core

Reduction of the Inhibition Power, % 4.3 10.3 14.6 1.9 3.3 7.9 13.1 12.9 12.4 2.6 4.0 3.9

tion of the antioxidant power from a minimum of 4.32% to a maximum of 14.60% for the higher antioxidant concentration (increasing trend). The same increasing trend was observed for linalool emulsions entrapped in the inner core; but, the average inhibition power reduction was lower than lipidic layer, from a minimum of 1.93% to a maximum of 7.90% for the highest antioxidant concentration. Similarly, limonene entrapped into lipidic layer showed an almost stable behavior by increasing antioxidant concentration, from 12.4% to 13.1%; whereas, emulsion entrapment guaranteed an inhibition reduction from 2.6% to 4.0%, increasing drug concentration. A general comment on these results could be that, since the compounds entrapped in the lipidic layer are more directly in contact with the expanded liquid, the effect of the high pressures and temperatures of 40 ◦ C could cause degradation phenomena. Instead, the entrapment in emulsions loaded in the inner core of liposomes better preserved the compounds, during the production and the following collection. 4. Conclusions In this work, a new approach for the inclusion of lipophilic drugs into liposome aqueous core was successfully proposed, by first stabilize the lipophilic molecule into the disperse phase of an oil in water emulsion. The emulsion was, then, entrapped into liposome inner core using SuperLip process. Results showed a favorable effect of supercritical emulsion processing on the re-arrangement of O/W emulsion droplets that lead to the production of a stable emulsion at nanometric level. Furthermore, encapsulation efficiencies were significantly increased using the emulsion inclusion strategy, up to 92% for limonene and 99% for linalool. The results obtained confirmed that water droplets atomized in the formation vessel can be considered discrete volumes in which the dissolved or suspended molecules are confined during the encapsulation process, resulting in higher entrapment efficiencies. The generalized result obtained from this research could represent a novel approach to the entrapment of lipophilic compounds into liposomes using oil in water emulsions.

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Acknowledgements The authors would like to acknowledge Paola Russo and Alessandra Saturnino for their bachelor degree work related to this projects. References [1] G.M. Barratt, Therapeutic applications of colloidal drug carriers, Pharm. Sci. Technol. Today (2000), http://dx.doi.org/10.1016/S1461-5347(00)00255-8. [2] A. Laouini, C. Jaafar-Maalej, I. Limayem-Blouza, S. Sfar, C. Charcosset, H. Fessi, Preparation, characterization and applications of liposomes: state of the art, J. Colloid Sci. Biotechnol. (2012), http://dx.doi.org/10.1166/jcsb.2012.1020. [3] A. Wagner, K. Vorauer-Uhl, Liposome technology for industrial purposes, J. Drug Deliv. (2011), http://dx.doi.org/10.1155/2011/591325. [4] J. Li, X. Wang, T. Zhang, C. Wang, Z. Huang, X. Luo, Y. Deng, A review on phospholipids and their main applications in drug delivery systems, Asian J. Pharm. Sci. (2014), http://dx.doi.org/10.1016/j.ajps.2014.09.004. [5] A. Samad, Y. Sultana, M. Aqil, Liposomal drug delivery systems: an update review, Curr. Drug Deliv. (2007), http://dx.doi.org/10.2174/ 156720107782151269. [6] M.R. Mozafari, C. Johnson, S. Hatziantoniou, C. Demetzos, Nanoliposomes and their applications in food nanotechnology, J. Liposome Res. (2008), http://dx. doi.org/10.1080/08982100802465941. [7] H.R. Ahmadi Ashtiani, P. Bishe, N.-A. Lashgari, M.A. Nilforoushzadeh, S. Zare, Liposomes in cosmetics, J. Ski. Stem Cell. (2018), http://dx.doi.org/10.5812/ jssc.65815. [8] L. Sercombe, T. Veerati, F. Moheimani, S.Y. Wu, A.K. Sood, S. Hua, Advances and challenges of liposome assisted drug delivery, Front. Pharmacol. (2015), http://dx.doi.org/10.3389/fphar.2015.00286. [9] H. Barani, M. Montazer, A review on applications of liposomes in textile processing, J. Liposome Res. (2008), http://dx.doi.org/10.1080/ 08982100802354665. [10] M. Alavi, N. Karimi, M. Safaei, Application of various types of liposomes in drug delivery systems, Adv. Pharm. Bull. (2017), http://dx.doi.org/10.15171/ apb.2017.002. [11] M. C¸a˘gdas¸, A.D. Sezer, S. Bucak, Liposomes as potential drug carrier systems for drug delivery, Appl. Nanotechnol. Drug Deliv. (2014), http://dx.doi.org/10. 5772/58459. [12] N. Maurer, D.B. Fenske, P.R. Cullis, Developments in liposomal drug delivery systems, Expert Opin. Biol. Ther. (2005), http://dx.doi.org/10.1517/14712598. 1.6.923. [13] S.K. Sahoo, F. Dilnawaz, S. Krishnakumar, Nanotechnology in ocular drug delivery, Drug Discov. Today (2008), http://dx.doi.org/10.1016/j.drudis.2007. 10.021. [14] N. Mignet, J. Seguin, G.G. Chabot, Bioavailability of polyphenol liposomes: a challenge ahead, Pharmaceutics (2013), http://dx.doi.org/10.3390/ pharmaceutics5030457. [15] N. Düzgünes¸, G. Gregoriadis, Introduction: The origins of liposomes: Alec Bangham at Babraham, Methods Enzymol. (2005), http://dx.doi.org/10.1016/ S0076-6879(05)91029-X. [16] E. Arab-Tehrany, B. Maherani, M.R. Mozafari, C. Gaiani, M. Linder, Liposomes: A Review of Manufacturing Techniques and Targeting Strategies, Curr. Nanosci. (2011), http://dx.doi.org/10.2174/157341311795542453. [17] M. Pons, M. Foradada, J. Estelrich, Liposomes obtained by the ethanol injection method, Int. J. Pharm. (1993), http://dx.doi.org/10.1016/03785173(93)90389-W. [18] H. Zhang, Thin-film hydration followed by extrusion method for liposome preparation, Methods Mol. Biol. (2017), http://dx.doi.org/10.1007/978-14939-6591-5 2. ´ I.T. Kostic, ´ A. Zvonar, V.B. Dordevic, ´ M. Gaˇsperlin, V.A. Nedovic, ´ [19] B.D. Isailovic, B.M. Bugarski, Resveratrol loaded liposomes produced by different techniques, Innov. Food Sci. Emerg. Technol. (2013), http://dx.doi.org/10. 1016/j.ifset.2013.03.006. [20] I. Espirito Santo, R. Campardelli, E.C. Albuquerque, S.A.B. Vieira De Melo, E. Reverchon, G. Della Porta, Liposomes size engineering by combination of ethanol injection and supercritical processing, J. Pharm. Sci. (2015), http://dx. doi.org/10.1002/jps.24595. [21] L. Lesoin, C. Crampon, O. Boutin, E. Badens, Preparation of liposomes using the supercritical anti-solvent (SAS) process and comparison with a conventional method, J. Supercrit. Fluids (2011), http://dx.doi.org/10.1016/j.supflu.2011.01. 006. [22] T. Imura, T. Gotoh, K. Otake, S. Yoda, Y. Takebayashi, S. Yokoyama, H. Takebayashi, H. Sakai, M. Yuasa, M. Abe, Control of physicochemical properties of liposomes using a supercritical reverse phase evaporation method, Langmuir. (2003), http://dx.doi.org/10.1021/la020589a. [23] L.A. Meure, R. Knott, N.R. Foster, F. Dehghani, The depressurization of an expanded solution into aqueous media for the bulk production of liposomes, Langmuir. (2009), http://dx.doi.org/10.1021/la802511a. [24] J.R. Falconer, D. Svirskis, A.A. Adil, Z. Wu, Supercritical fluid technologies to fabricate proliposomes, J. Pharm. Pharm. Sci. (2015). [25] R. Campardelli, P. Trucillo, E. Reverchon, Supercritical assisted process for the efficient production of liposomes containing antibiotics for ocular delivery, J. CO2 Util. (2018), http://dx.doi.org/10.1016/j.jcou.2018.04.006.

[26] P. Trucillo, R. Campardelli, E. Reverchon, Encapsulation of hydrophilic and lipophilic compounds in Nanosomes Produced with a supercritical based process, Lect. Notes Bioeng. (2018), http://dx.doi.org/10.1007/978-3-31962027-5 3. [27] R. Campardelli, P. Trucillo, E. Reverchon, Supercritical assisted process for the efficient production of liposomes containing antibiotics for ocular delivery, J. CO2 Util. (2018), http://dx.doi.org/10.1016/j.jcou.2018.04.006. [28] P. Trucillo, R. Campardelli, E. Reverchon, Production of liposomes loaded with antioxidants using a supercritical CO2assisted process, Powder Technol. (2018), http://dx.doi.org/10.1016/j.powtec.2017.10.007. [29] F. Shahidi, Y. Zhong, Measurement of antioxidant activity, J. Funct. Foods (2015), http://dx.doi.org/10.1016/j.jff.2015.01.047. [30] C¸. Yücel, G. S¸eker-Karatoprak, Development and evaluation of the antioxidant activity of liposomes and nanospheres containing rosmarinic acid, Farmacia (2017). [31] A.M. Galvão, J.S. Galvão, M.A. Pereira, P.G. Cadena, N.S.S. Magalhães, J.B. Fink, A.D. de Andrade, C.M.M.B. de Castro, M.B. de Sousa Maia, Cationic liposomes containing antioxidants reduces pulmonary injury in experimental model of sepsis: liposomes antioxidants reduces pulmonary damage, Respir. Physiol. Neurobiol. (2016), http://dx.doi.org/10.1016/j.resp.2016.06.001. [32] R. Gharib, H. Greige-Gerges, S. Fourmentin, C. Charcosset, L. Auezova, Liposomes incorporating cyclodextrin-drug inclusion complexes: current state of knowledge, Carbohydr. Polym. (2015), http://dx.doi.org/10.1016/j. carbpol.2015.04.048. [33] Z. Wen, B. Liu, Z. Zheng, X. You, Y. Pu, Q. Li, Preparation of liposomes entrapping essential oil from Atractylodes macrocephala Koidz by modified RESS technique, Chem. Eng. Res. Des. (2010), http://dx.doi.org/10.1016/j. cherd.2010.01.020. [34] H. Tavakoli, O. Hosseini, S.M. Jafari, I. Katouzian, Evaluation of physicochemical and antioxidant properties of yogurt enriched by olive leaf phenolics within Nanoliposomes, J. Agric. Food Chem. (2018), http://dx.doi. org/10.1021/acs.jafc.8b02759. [35] A. Rezaei, M. Fathi, S.M. Jafari, Nanoencapsulation of hydrophobic and low-soluble food bioactive compounds within different nanocarriers, Food Hydrocoll. (2019), http://dx.doi.org/10.1016/j.foodhyd.2018.10.003. [36] T. Ghorbanzade, S.M. Jafari, S. Akhavan, R. Hadavi, Nano-encapsulation of fish oil in nano-liposomes and its application in fortification of yogurt, Food Chem. (2017), http://dx.doi.org/10.1016/j.foodchem.2016.08.022. [37] A. Faridi Esfanjani, E. Assadpour, S.M. Jafari, Improving the bioavailability of phenolic compounds by loading them within lipid-based nanocarriers, Trends Food Sci. Technol. (2018), http://dx.doi.org/10.1016/j.tifs.2018.04.002. [38] P. Trucillo, R. Campardelli, M. Scognamiglio, E. Reverchon, Control of liposomes diameter at micrometric and nanometric level using a supercritical assisted technique, J. CO2 Util. (2019), http://dx.doi.org/10.1016/j.jcou.2019. 04.014. [39] P. Trucillo, R. Campardelli, E. Reverchon, Supercritical CO2 assisted liposomes formation: optimization of the lipidic layer for an efficient hydrophilic drug loading, J. CO2 Util. 18 (2017) 181–188, http://dx.doi.org/10.1016/j.jcou.2017. 02.001. [40] R. Campardelli, P. Trucillo, E. Reverchon, A Supercritical Fluid-Based Process for the Production of Fluorescein-Loaded Liposomes, Ind. Eng. Chem. Res. (2016), http://dx.doi.org/10.1021/acs.iecr.5b04885. [41] P. Trucillo, R. Campardelli, E. Reverchon, Operative parameters optimization production of liposomes for the encapsulation of hydrophilic compounds using a new supercritical process, Chem. Eng. Trans. (2017), http://dx.doi.org/ 10.3303/CET1757134. [42] R. Campardelli, P. Trucillo, E. Reverchon, Supercritical assisted process for the efficient production of liposomes containing antibiotics for ocular delivery, J. CO2 Util. 25 (2018) 235–241, http://dx.doi.org/10.1016/j.jcou.2018.04.006. [43] P. Panwar, B. Pandey, P.C. Lakhera, K.P. Singh, Preparation, characterization, and in vitro release study of albendazole-encapsulated nanosize liposomes, Int. J. Nanomedicine (2010). [44] S.J. Wallace, J. Li, R.L. Nation, B.J. Boyd, Drug release from nanomedicines: selection of appropriate encapsulation and release methodology, Drug Deliv, Transl. Res. (2012), http://dx.doi.org/10.1007/s13346-012-0064-4. [45] M.G. Miguel, Antioxidant and anti-inflammatory activities of essential oils: a short review, Molecules. (2010), http://dx.doi.org/10.3390/ molecules15129252. [46] W. Dhifi, S. Bellili, S. Jazi, N. Bahloul, W. Mnif, Essential oils’ chemical characterization and investigation of some biological activities: a critical review, Medicines (2016), http://dx.doi.org/10.3390/medicines3040025. [47] M. Aissou, Z. Chemat-Djenni, E. Yara-Varón, A.S. Fabiano-Tixier, F. Chemat, Limonene as an agro-chemical building block for the synthesis and extraction of bioactive compounds, Comptes Rendus Chim. (2017), http://dx.doi.org/10. 1016/j.crci.2016.05.018. [48] A.M.J.C. Neto, G.V.S. Mota, R.S. Borges, M.L.S. Albuquerque, R-(-)linalool UV spectroscopy: the experimental and theoretical study, J. Comput. Theor. Nanosci. (2010), http://dx.doi.org/10.1166/jctn.2010.1375. [49] J.M. RANDALL, W.L. BRYAN, O.W. BlSSETT, R.E. BERRY, Ultraviolet Absorption Method for evaluating Citrus Essences, J. Food Sci. (1973), http://dx.doi.org/ 10.1111/j.1365-2621.1973.tb02145.x. [50] R.G. Bostedor, J.D. Karkas, B.H. Arison, V.S. Bansal, S. Vaidya, J.I. Germershausen, M.M. Kurtz, J.D. Bergstrom, Farnesol-derived dicarboxylic acids in the urine of animals treated with zaragozic acid A or with farnesol, J. Biol. Chem. (1997), http://dx.doi.org/10.1074/jbc.272.14.9197.

P. Trucillo, R. Campardelli and E. Reverchon / J. of Supercritical Fluids 154 (2019) 104626 [51] W. Brand-Williams, M.E. Cuvelier, C. Berset, Use of a free radical method to evaluate antioxidant activity, LWT - Food Sci. Technol. (1995), http://dx.doi. org/10.1016/S0023-6438(95)80008-5. [52] V.B. Henriques, R. Germano, M.T. Lamy, M.N. Tamashiro, Phase transitions and spatially ordered counterion association in ionic-lipid membranes: theory versus experiment, Langmuir (2011), http://dx.doi.org/10.1021/ la202302x. [53] M. Ochowiak, The effervescent atomization of oil-in-water emulsions, Chem. Eng. Process. Process Intensif. (2012), http://dx.doi.org/10.1016/j.cep.2011.11. 007. [54] J.M. Perrier-Cornet, P. Marie, P. Gervais, Comparison of emulsification efficiency of protein-stabilized oil-in-water emulsions using jet, high pressure

9

and colloid mill homogenization, J. Food Eng. (2005), http://dx.doi.org/10. 1016/j.jfoodeng.2004.03.008. [55] J. Floury, A. Desrumaux, J. Lardières, Effect of high-pressure homogenization on droplet size distributions and rheological properties of model oil-in-water emulsions, Innov. Food Sci. Emerg. Technol. (2000), http://dx.doi.org/10.1016/ S1466-8564(00)00012-6. [56] H. Karbstein, H. Schubert, Developments in the continuous mechanical production of oil-in-water macro-emulsions, Chem. Eng. Process. Process Intensif. (1995), http://dx.doi.org/10.1016/0255-2701(94)04005-2. [57] P. Marie, J.M. Perrier-Cornet, P. Gervais, Influence of major parameters in emulsification mechanisms using a high-pressure jet, J. Food Eng. (2002), http://dx.doi.org/10.1016/S0260-8774(01)00138-8.