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Sonication technique to produce emulsions: The impact of ultrasonic power and gelatin concentration
Accepted Manuscript Sonication technique to produce emulsions: The impact of ultrasonic power and gelatin concentration Karen Cristina Guedes Silva, Ana Carla Kawazoe Sato PII: DOI: Reference:
S1350-4177(18)31159-3 https://doi.org/10.1016/j.ultsonch.2018.12.001 ULTSON 4398
To appear in:
Ultrasonics Sonochemistry
Received Date: Revised Date: Accepted Date:
1 August 2018 30 November 2018 1 December 2018
Please cite this article as: K.C.G. Silva, A.C.K. Sato, Sonication technique to produce emulsions: The impact of ultrasonic power and gelatin concentration, Ultrasonics Sonochemistry (2018), doi: https://doi.org/10.1016/ j.ultsonch.2018.12.001
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Sonication technique to produce emulsions: the impact of ultrasonic power and gelatin concentration Karen Cristina Guedes Silva a1, Ana Carla Kawazoe Satoa2. a
Department of Food Engineering, School of Food Engineering (FEA), University of
Campinas, CEP: 13083-862, Campinas, SP, Brazil. 1
[email protected], [email protected].
Corresponding authors: Department of Food Engineering, School of Food Engineering (FEA), University of Campinas. Rua Monteiro Lobato, 80, CEP: 13083-862, Campinas, SP, Brazil. Tel.: +55 19 35214088. E-mail addresses: [email protected] (K. C. G. Silva), [email protected] (A. C. K. Sato).
Highlights • Oil in water emulsions was prepared by rotor-stator and sonication processes. • Sonication process reduces the viscosity of the aqueous phase. • Higher ultrasonic power produced emulsions with higher stability. • Low protein concentration led to a coalescence during sonication. • Sonication was proposed as a suitable method for emulsion production.
ABSTRACT The production of food emulsions has increased the demand for processes, natural emulsifiers and stabilizers that provide reasonable stability. This study approaches the influence of parameters that affect the stability of emulsions produced by sonication, such as ultrasonic power (150, 225 and 375 W) and gelatin concentration, when producing alginate, potato starch and gelatin stabilized emulsions. The results showed that sonication reduced viscosity, surface charge and improved the interfacial properties of biopolymeric solutions. Emulsions presented visual kinetic stabilization after 7 days of storage. The increase of sonication power reduced particle size but increased flocculation. The use of ultrasonic power at 225 and 375W and gelatin in a concentration above 1.0 % resulted in stable emulsions with smaller particle size, which is desirable for its application in food systems.
Keywords: ultrasound, alginate, potato starch, emulsification, aqueous phase, stability.
1.
Introduction Emulsions are mixtures of immiscible liquids containing droplets of one liquid
(dispersed phase) dispersed in a continuous liquid phase. Emulsifying agents are the ones that provide stability to these systems. However, the emulsification process is entropically unfavorable, resulting in thermodynamically unstable systems [1]. Hence, it is important to develop highly stable food emulsions with no or minimal changes in its structure during its process and storage [2]. Different processes have been studied to achieve greater emulsion stability: high energy processes such as rotor-stator, ultrasound devices, high-pressure homogenizer, microfluidics, among others widely employed [3,4]. Previous studies have shown that, for high energy processes, the type of homogenization process, operating conditions (energy intensity, time and temperature), sample composition (oil and emulsifier concentrations), and physicochemical properties of the compounds (interfacial tension and viscosity), result in emulsions with different characteristics [5]. Commonly used to prepare pre-emulsions, rotor-stator process mixes liquids from separated phases, forcing them through a narrow gap between a rotating disc (rotor) and a static disk (stator). The rapid flow circulation and high shear within the narrow gap between the rotor and stator form droplets usually larger than those obtained
by other high energy processes [4,6]. It is possible to produce finer emulsions by sonication. In this process, the mechanism for droplet rupture is acoustic cavitation, which comprises the growth and collapse of micro gas bubbles dissolved in the medium, leading to droplets breakdown [7]. One can apply the ultrasonic devices to liquid media in the form of bath or probe. In ultrasonic baths, the transducers are evenly located at the bottom of the bath. However, interferences between waves can result in different wave intensity at different positions of the tank [8]. The probe system consists of a generator and a converter, which transforms energy into mechanical vibrations transmitted by a probe and directly applied to the sample afterwards [4]. In general, probe type sonicators have higher localized intensity and effect compared to those of bath type [9]. When compared to homogenizers and microfluidizers, sonication processes feature advantages such as being easier to clean and presenting a simpler system, not needing external pressure, using less energy and using less surfactant to obtain more homogenous emulsions [10,11]. With ultrasound application, benefits have been obtained in food processing and preservation, such as higher product yields, shorter processing times and lower costs, in addition to an environmentally friendly production [12]. Moreover, structural modifications from sonication of food structures enable the development of new products with unique properties and quality attributes [13]. In this sense, sonication can lead to functional modifications in polymer chains [14,15] and protein structures [16,17], by adding functional features, such as improvements in dissolution, hydration, hydrophobicity, emulsifying and rheological performances [16], besides improving antioxidant properties of certain food materials [18]. The ultrasound application has also shown to modify some physical properties of starch and biopolymers solutions, allowing its different applications [19–21]. Besides the process, biopolymers used to stabilize emulsions play a fundamental role in stabilization. Some papers in the literature have shown that the association of proteins and polysaccharides for emulsion production can increase stability during the process, compared to emulsions only stabilized by proteins [22]. This association contributes to the structure, texture and stability of food systems, due to the thickening or gelling behavior and surface properties of biopolymers [23,24]. Few studies provide a systematic approach to evaluate the effects of the ultrasound process on emulsion formation and stabilization, often associated with beneficial structural modifications of biopolymers. These modifications, allied with the
establishment of ideal concentrations of surfactant, structuring agents and energy employed in the process, may result in emulsions with improved characteristics and greater stability. Therefore, the objective of this study was to investigate the effects of the ultrasound process on the physical properties of natural biopolymers used to stabilize emulsions, such as protein and polysaccharides, and to understand how the ultrasound-promoted structural modifications of biopolymers can influence emulsion stability. Hence, it would be possible to apply these emulsions to food systems of varied characteristics as structuring agents, or even as encapsulation systems.
2. Materials and methods 2.1. Materials To prepare the emulsions, ingredients such as alginate FD 17X, donated by Danisco (France), A-type gelatin - bloom 280 (~5.5 isoelectric point and 87.92 g/100 ml protein content), supplied by Gelco (Brazil), potato starch from Zona Cerealista (Brazil) and sunflower oil, purchased from a local supermarket, were used. All other chemicals were of analytical grade or equivalent.
2.2. Methods 2.2.1. Aqueous phase preparation Solutions containing a mixture of 3% (w/w) potato starch, 0.5% (w/w) alginate and varying amounts of gelatin (0, 0.1, 0.5, 1.0 and 1.5 % (w/w)), named as SA, SAG0.1 ; SAG0.5 ; SAG1.0 and SAG1.5, respectively, were prepared. Solutions presented pH 6 ± 0.2. Starch and alginate were added into deionized water at room temperature, under constant magnetic stirring, until the alginate was dissolved and no lumps were observed. Subsequently, solutions were heated at 90 °C under constant magnetic stirring at 800 rpm, for 20 min. The solution was cooled to up to 45 °C before the addition of different concentrations of gelatin, maintaining constant magnetic stirring at 800 rpm for 20 min. Rheology of biopolymer solutions was evaluated. Pure starch (S), Alginate (A) and Gelatin (G) solutions were prepared at the same concentrations mentioned above, under the same conditions. These solutions were used as a control to evaluate the influence of ultrasound on the properties of each biopolymer separately.
2.2.2. Emulsion preparation
Oil-in-water emulsions were prepared by mixing 20% (w/w) sunflower oil and 80% (w/w) of aqueous phase solution, using a rotor-stator (RS) device (SilentCrusher M, Heidolph, Germany) at 20000 rpm for 4 min prior to bath or probe sonication. For the ultrasonic probe, an ultrasonic processor (QR 750 W, Ultronique, Brazil) with a frequency of 20 kHz, attached to a titanium probe (13 mm diameter) was used to sonicate the emulsions. Sonication time was fixed at 5 min, its power varying in 150 (UP1), 225 (UP2) and 375 W (UP3). A glass tube containing 30 mL of sample was introduced in jacketed beakers at 10 oC to dissipate the heat produced during sonication and keep the temperature below 45 oC. Ultrasonic bath (USC 1450, Unique, Brazil) was tested with a power of 150 W and 25 kHz frequency, and sonication time fixed at 5 min. Samples submitted to the ultrasonic bath treatment were named as UB. Emulsions were evaluated regarding their stability, microscopy and particle size. Sonication time was set at 5 min for bath and probe sonication equipment, once preliminary tests showed that increasing sonication time resulted in larger droplets (Table 1), indicating an over processing, possible due to a higher probability of droplet collision, resulting in coalescence [25].
2.3. Biopolymer solutions characterization 2.3.1. Differential Scanning Calorimetry Thermal properties of starch were analyzed using a scanning differential calorimeter (DSC 2920, TA Instruments, USA). The sample was heated at a 10 °C/min rate from 10 to 110 °C. The initial temperature (To), peak temperature (Tp), final temperature (Tf), and gelatinization enthalpy (ΔH) were obtained from sample thermograms analyzed using the Universal Analysis software (TA Instruments). 2.3.2. ζ - potential ζ-potential of biopolymers was determined using a Zetasizer Nano Series (Malvern Instruments, UK). Biopolymers solutions were measured with and without ultrasound application at the same concentration prepared for the aqueous phase.
2.3.3. Interfacial tension Interfacial tension between biopolymer solutions in water and sunflower oil was evaluated using a tensiometer (Tracker-S, Teclis, France) by the pendant (O/W) drop method. Interfacial properties of gelatin at different concentrations and the effects of
ultrasound on the interfacial properties of alginate, gelatin and starch was evaluated. The test was conducted for over 7200 s and the temperature was maintained at 25 °C throughout the test.
2.3.4. Rheological measurements Rheological properties of aqueous phase were characterized using a rheometer (AR1500ex, TA Instruments, USA) equipped with a cone/plate geometry (cone diameter: 40 mm; angle: 1,58° and gap size: 53 μm). Flow curves were obtained in the shear rate range of 0-1000 s-1 at 25 °C. Experimental data were described by the power law equation (1): ( 1) where and
is shear stress (Pa),
is consistency index (Pa. sn),
is shear rate (s-1)
is the flow behavior index (-).
2.4. Emulsions evaluation 2.4.1. Stability Immediately after preparation, 25 ml of each emulsion was poured into a cylindrical glass tube, sealed with a plastic cap and stored in BOD (MA 415 UR, Marconi, Brazil) at 25 °C for 7 days. The O/W emulsion stability was measured by the height of the bottom phase (Hb) and expressed according to creaming index (CI) (equation 2). ( (2) where H0 represents the initial height of emulsion. The analyses were carried out in duplicate.
2.4.2. Size distribution Measurement of droplet size distribution was performed in a Malvern Mastersizer S (Malvern Instruments, UK) equipped with accessory Hydro 2000S (Malvern Instruments, UK). Samples were dispersed into distilled water. Values of Sauter mean diameter, which is inversely proportional to the specific surface area of
droplets, were obtained (Equation 3). The polydispersity (Span) was also evaluated (Equation 4). n
D3, 2
n.d
3 i
n.d
2 i
i 1 n
i 1
( 3)
( 4) where ni is number of droplets with a diameter di, and
(µm) are
diameters at 10, 50 and 90% of cumulative volume, respectively.
2.4.3. Optical microscopy Emulsion microstructure was observed in an optical microscope (Axio Scope.A1, Carl Zeiss, Germany) with a 100× oil immersion objective lens. Images were captured with the software AxioVision Rel. 4.8 (Carl Zeiss, Germany). Optical microscopy was performed in freshly prepared emulsions and after 7 days of storage. The public domain software Image J v1.36b (http://rsb.info.nih.gov/ij/) was used to count droplets and measure their size.
2.6. Statistical analyses All trials were replicated three times and, unless otherwise stated, all analyses were carried out in two independent replicates. SISVAR software - Computer Statistical Analysis System [26] was used for variance analysis (ANOVA). Significant differences (p<0.05) between samples were determined by the Tukey procedure.
3. Results 3.1. Effect of gelatin concentration and process at the aqueous phase Aiming to evaluate the role of protein at different concentrations of emulsion stabilization, in addition to the effects of mechanical and sonication process in the structure of the isolated biopolymers, surface charge analysis, interfacial tension and rheology were performed for aqueous phase.
3.1.1. Differential Scanning Calorimetry
To determine the gelatinization temperature of the starch, we measured its thermal properties: T0 = 62.29 ± 0.11 °C, Tp = 65.64 ± 0.38 °C, Tf = 70.12 ± 0.11 °C and gelatinization enthalpy (∆H) (8.93 ± 1.35 J/g dry starch). The starch dilutions employed in this study were higher than 10 g water/g potato starch for all formulations, which ensures complete gelatinization, as the gelatinization temperature of the experiments was set at 90 °C/20 min. Evans & Hais [27] reported values for initial (~60 °C) and final (~70 °C) gelatinization temperature as a function of water content, for a ratio higher than 2 g water/g potato starch. 3.1.2. ζ - potential Zeta potential results of biopolymers showed that the sonication process (UP3) significantly reduced surface charge for all biopolymers (p<0.05) (Tab. 2).
3.1.3. Interfacial tension The evaluation of interfacial tension was carried out with the objective to characterize the interfacial properties of solutions with different gelatin concentrations and to investigate the influence of the sonication process on the emulsifying properties of biopolymers. Regarding interfacial properties of the protein, the increase of gelatin concentration exerts little influence on initial interfacial tension (7.2 – 7.8 mN/m). The difference became more pronounced at 7200 s, presenting results that varied between 3.1 to 4.3 mN/m for G1.5 to G0.1, respectively (Fig. 1A), which may be related to faster adsorption of gelatin at the droplet interface. Higher adsorption of the surfactant at the interface is associated with a reduction in interfacial properties, indicating that the surfactant and its amount have been effective, forming a protective barrier that prevents flocculation and retards destabilization [3]. However, it should be noted that even though higher concentration of surfactant reduces interfacial tension, the equilibrium in interfacial tension was not reached after 7200 s. Concerning the influence of the ultrasonic process (UP3), a decrease in the interfacial tension for gelatin and alginate solutions was observed compared to the same solutions without treatment. One can associate this result with the exposure of hydrophobic residues of the surfactant during sonication [28], which is also possible to observe from surface charge reduction after ultrasound application (Tab. 2). In contrast,
starch solution increased in interfacial tension when submitted to sonication process (UP3) (Fig. 1B).
3.1.4. Rheological measurements The ultrasound process employs a large amount of energy generating hydrodynamic shear forces, causing cavitation and collapse of gas bubbles. This phenomenon can change physical or physicochemical structures of the medium [16,29], which may result in the breaking of some chains, allowing greater fluidity of the continuous phase. Rheology results for isolated biopolymer solutions showed that sonication process (UP3) reduced the viscosity of alginate, gelatin and starch, which was of about two orders of magnitude for the latter (Tab. 3). Sonication can accelerate the depolymerization of starch gels, separating amylose and amylopectin units. In this situation, amylose units remain in the gel amorphous phase, which leads to the weakening of biopolymer network, thus reducing viscosity [19]. However, authors point out that despite sonication dramatically affecting solution viscosity, the chemical structure of polymer chains may not change [19]. The effect of process on the rheological properties of aqueous phase containing the associated biopolymers (Tab. 4) showed that rheological parameters of solutions submitted to probe sonication (UP) differed significantly (p<0.05) from the same formulation submitted to bath sonication (UB) or rotor stator (RS). The increase in sonication power reduced shear thinning of SAG solutions (n closer to 1) and reduced viscosity (associated to lower k) (Tab. 4). This effect may be due to possible breakdown of polymer chains caused by the sonication process [29,30]. Such breakdown reduces entanglements and consequently modifies intermolecular interactions between polymer molecules, reducing viscosity and leading to an approximation of Newtonian behavior [31,32]. Similar results regarding viscosity reduction with ultrasound application in polymers, such as glucomannan, pectin, proteins and starches from different sources have been reported in literature [19,32,33]. By comparing the same treatment through varying concentrations of gelatin, we observed that the increase in gelatin concentration increased viscosity for RS, UB and UP1. The effect of gelatin concentration is suppressed at higher powers (UP2 and UP3), with no significant differences (p>0.05) in viscosity (Tab. 4).
3.2. Effect of gelatin concentration and process on emulsions The power used during emulsification is a key point for the production of stable emulsions. At this stage of the work, we used different concentrations of gelatin when producing emulsions by stator rotor, ultrasonic bath and ultrasonic probe, at three different sonication powers. Stability, droplet size, and emulsion morphology were investigated.
3.2.1. Creaming index Emulsions were evaluated regarding stability by creaming index. Gelatin control emulsions submitted to RS, UB and UP1 process showed creaming index of 39.23 ± 3.39, 38.80 ± 2.06 and 24.71 % ± 6.60, respectively. For the same formulation, UP2 and UP3 process did not present phase separation at 7 days (Fig. 2A). The higher stability presented by emulsions solely containing gelatin was only observed for conditions of higher sonication power. This result supports the hypothesis that higher sonication power favors the exposure of surfactant hydrophobic residues, as fat-binding ability depends on the degree of exposure of gelatin hydrophobic residues [21] (Tab. 2 and Fig. 1B). For control emulsions without gelatin (SA), all formulations presented phase separation. The creaming index for RS, UB, UP1, UP2 and UP3 were: 60.92 ± 2.58, 61.82 ± 1.29, 66.73 ± 4.49, 43.97 ± 7.19 and 65.25 % ± 1.84, respectively (Fig. 2B). Despite sonication changing zeta potential, rheology and interfacial tension of alginate and starch solutions (Tab. 2 and 3 and Fig. 1B), neither of them act as emulsifiers, only contributing to system viscosity [23]. Thus, no stable emulsion was produced solely with alginate and starch. Protein association with potato starch and alginate (SAG0.1 to SAG1.5) provided visually stable emulsions under all tested processes and concentrations without creaming (data not shown), which emphasizes the importance of interfacial activity promoted by gelatin and of the hydrodynamic stabilization [3] promoted by starch and alginate.
3.2.2. Droplet size and morphology during storage Emulsions were evaluated in relation to their droplet size, polydispersity (SPAN), optical microscopy and image analysis at day 0 and after 7 days of storage (Tab. 5). Results showed that, for the same process, gelatin concentration (0.5 to 1.5%
(m/m)) did not show significant differences (p>0.05) (statistical data not shown) for emulsion droplet size. In other words, increase in sonication power was more effective in emulsion size reduction compared to gelatin concentration. Comparing the different processes at a fixed protein concentration, it was possible to verify the treatment influence on droplet size and, consequently, on stability. For the lowest protein concentration (SAG0.1) and for control emulsion (SA), we observed no significant differences (p>0.05) between RS, UB and UP1 treatments at day 0. However, the treatment with the highest sonication power (UP3) resulted in larger particle size immediately after the process (day 0). Higher sonication powers (UP2 and UP3) expose the droplets to large amounts of energy, not only led to droplet rupture but also collision and, therefore, coalescence and flocculation [34]. Possibly, only adding 0.1% gelatin was not sufficient to cover and stabilize the droplet interface [35]. In addition, viscosity reduction due to the process (Tab. 4) may have aggravated destabilization under these conditions [36]. During the 7 days of storage, destabilization became more pronounced, as can be observed in D[3,2] results (Tab. 5). Addition of 0.5 and 1.0% of protein to the emulsion resulted in increase of D[3,2] with increasing ultrasound power. However, size results obtained by image analysis indicated a reduction, as we can see in micrographs (Tab. 5). Such divergence between image analysis and laser diffraction (D[3,2] result), reflects flocculation caused by the process. The smallest droplet sizes of SAG0.5 and SAG1.0 came from the combination of gelatin concentration and ultrasound treatment. At these concentrations, it would be possible to coat the droplets formed during emulsification efficiently. Moreover, the most intense cavitation process (UP2 and UP3) reduced the viscosity of aqueous phase, facilitating surfactant mobility to the droplet interface, avoiding coalescence and reducing polydispersity [3]. Analyzing the results obtained for SAG1.5 emulsion, it was observed that droplet size also tends to reduce as ultrasonic power increases. However, SAG1.5 presented higher flocculation and apparent gelation at 7 days of storage (Tab. 5), despite being stored at room temperature (25 °C).
4. Discussion The use of higher sonication powers led to structural modifications in the aqueous phase of the isolated biopolymers, which reduced biopolymer viscosity (Table 3), interfacial tension (Figure 1), and surface charge intensity (Table 2). For gelatin,
these results are associated to the exposure of hydrophobic protein residues to sonication, reducing charge intensity [16]. We can associated such exposure with improvements in stability and dehydration of amphiphilic biopolymers, which decreases the energy required for protein adsorption at the oil-water interface [30]. The breakage of electrostatic and hydrophobic interactions between protein molecules with ultrasound has facilitated the hydrolysis process of gelatin [37] and increased solubility and thermal stability of treated whey protein[17]. Structural modifications provided by sonication has also been used to regulate viscosity [38] and modify foaming properties of products [13,39]. Structural modifications of several biopolymers have also been reported through sonication in the literature. Hosseini et al., [15] also reported the ζ-potential reduction for β-lactoglobulin and κ-carrageenan complexes with sonication. The authors attributed this phenomenon to the reduction of biopolymer reactivity after sonication, related to heterogeneous sonochemical interactions and structural changes. These changes can include a large number of sonochemical reactions in polysaccharides, including glycosylation, acetalization, oxidation and carbon bonds, capable of modifying interactions between polysaccharides and reducing reactive sites [12]. In this study, the zeta potential reduction observed for alginate and starch may stem from these sonochemical reactions. Viscosity reduction may depend on mechanical, structural properties of biopolymers, on linear or random coil configurations, as discussed by Iida et al., [19] for pectin and glucomannan. For individual biopolymers, we observed different intensity on viscosity reduction depending on their structure. Potato starch, the compound with the highest molecular weight [40], also showed the highest viscosity reduction compared to gelatin, a protein with random coil structure [41]. It is known that viscosity of the continuous phase directly affects the stability of dispersions [7]. By applying the highest powers (UP2 and UP3), we observed higher viscosity reduction at the aqueous phase. Such reduction provides greater surfactant mobility to interface droplets, which quickly reduces interfacial tension [42], as observed in this study. Behrend and Schubert [7] also reported that the higher viscosity of the continuous phase delays the mobility and stabilization of surfactant at the interface. Some studies have shown that the association of proteins and polysaccharides in emulsion production may increase the stability when compared to emulsions only stabilized by proteins [22]. The action of hydrocolloids as structuring, thickening and
gelling agents at the aqueous phase [43] provides a rheological control mechanism that immobilizes the oil droplets in an entangled biopolymer network [44]. In processes applying lower sonication powers (RS, UB and UP1) due to the higher viscosity, it is possible to say that part of the gelatin adhered to the surface of the droplets while part was dispersed in the medium with alginate and starch. This configuration characterizes depletion stabilization, a mechanism in which free (nonadsorbed) polymers inhibits aggregation [45], and in addition to the hydrodynamic stabilization from high viscosity. This behavior is often found in mixtures of food biopolymers [46]. Sonication at higher powers (UP2 and UP3) reduce droplet size and increased flocculation, which
is clear when observing the divergence between D[3,2] data,
obtained by laser diffraction, and size, obtained by optical microscopy images (Tab. 5). Under these conditions, it is possible to consider that the sonicated protein in solution has migrated to the oil interface, reducing the protein/polysaccharide ratio in the medium. It is worth noting that the energy employed in the ultrasound process is not sufficient to break the peptide bonds, but only non-covalent associative interactions, which does not cause changes in the primary structure of proteins [16]. Increases in free polysaccharide solution favor depletion flocculation [47, 48]. In this study, the presence of non-adsorbed alginate and starch in the continuous phase, which has no interfacial properties caused an osmotic effect, reducing the amount of solvent between the droplets and promoting attraction between them. This phenomenon occurs when protein-coated droplets are dispersed in an aqueous solution of other biopolymer not compatible with the protein [49]. In this case, the repulsion between gelatin located at the interface with other biopolymers occurs because all of them present a negative surface charge under these study conditions (Tab. 2). The increase in the size of bovine gelatin chains after 7 days of storage was also reported, indicating a reorganization of proteins in sub-aggregates due to non-covalent (electrostatic and hydrophobic) interactions [30]. This reorganization may have been responsible for the increased flocculation observed for the emulsions obtained with the highest intensity processes (UP2 and UP3) after 7 days (Tab. 5). The role of ultrasound in protein functionality modifications, such as thermal, structural and morphological attributes [17], and nanoemulsion production functionality has been studied [16]. However, few works in the literature have studied the effects of ultrasound process on structural modifications of the separate biopolymers and its
effects when biopolymers are associated, trying to understand how these modifications contribute to stabilizing emulsions. Besides, there has been increasing interest within the food industry to replace synthetic emulsifiers by natural ones, as an alternative to consumer-friendly products. Consequently, the trend of using biopolymers (proteins, starches) to produce and stabilize emulsions is growing [50] and there is still need to further understand the functionality of these biopolymers, when applied to different emulsification process. The results of this study indicated that the type of process and its intensity caused structural modifications in biopolymers. These modifications influenced the characteristics of the aqueous phase and consequently the formation of emulsions, resulting in systems with different characteristics.
4. Conclusions This study showed that the type of process provided more emulsion modifications, resulting in a wider variety of emulsions when compared to gelatin concentration. Higher sonication power changed the polymer network, with more influence on starch. Generally, sonication process decreased zeta potential, interfacial tension and viscosity of the biopolymers solutions, resulting in stable emulsions with smaller droplet size. Emulsions SAG0.5 and SAG1.0 sonicated at 225 and 375 W presented desirable characteristics for food application. Besides using only natural compounds, this study demonstrated that producing emulsions by sonication employing higher powers (UP2 and UP3) is a simple process that promotes desirable structural modifications in biopolymers.
Acknowledgments The authors thank the access to equipment and assistance provided by the laboratory of Process Engineering, Department of Food Engineering, School of Food Engineering from the State University of Campinas co-funded by Fapesp (2007/580175; 2009/54137-1; 2011/06083-0; 2015/11984-7). Karen Cristina Guedes Silva thanks CAPES for the assistantship. The authors thank Espaço da Escrita – Pró-Reitoria de Pesquisa - UNICAMP - for the language services provided.
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Fig. 1: (A) Interfacial tension of sunflower oil in gelatin solution at concentrations 0.1 % (w/w), 0.5 % (w/w), 1.0 % (w/w) and 1.5 % (w/w). (B) Interfacial tension of sunflower oil in gelatin 1.0 % (w/w), alginate 0.5 % (w/w) and starch 3.0 % (w/w) solutions. Solutions without treatment and submitted to ultrasonic power at 375 W/5 min (UP3) using the pendant drop method. Fig. 2: Stability of control emulsions at 7 days, submitted to rotor stator (RS), ultrasonic bath at 150 W/5 min (UB), ultrasonic probe at 150 W/5 min (UP1), 225 W/5 min (UP2) and 375 W/5 min (UP3). (A) Emulsion of sunflower oil in gelatin solution 1.5% (w/w). (B) Emulsion of sunflower oil in starch 3.0% (w/w) and alginate 0.5% (w/w) solutions.
A
B
G RS
G UB
G UP1
G UP2
G UP3
SA RS
SA UB
SA UP1
SA UP2
SA UP3
Tab. 1: Polydispersity (Span) and Sauter mean diameter (D3,2) of the SAG1.0 emulsion (0 days) submitted to ultrasonic bath (UB) and ultrasonic probe at 375 W (UP3) at 5, 10 and 15 min. Formulations SAG1.0 UB 5min SAG1.0 UB 10min SAG1.0 UB 15min SAG1.0 UP3 5min SAG1.0 UP3 10min SAG1.0 UP3 15min
Span 1.637 ± 0.002b 1.673 ± 0.012b 1.865 ± 0.008c 1.484 ± 0.010a 1.925 ± 0.005d 1.923 ± 0.012d
Lowercase letters show differences (p < 0.05) between the formulations
D [3, 2] (µm) 2.547 ± 0.002a 2.399 ± 0.028a 2.762 ± 0.077b 4.216 ± 0.140c 5.541 ± 0.082d 6.054 ± 0.040e
Tab. 2: Zeta potential of biopolymers alginate (A), gelatin (G) and potato starch (S) at pH 6 ± 0.2. Solutions without treatment and submitted to ultrasonic process at 375 W/5 min (UP3). Formulations A0.5 G1.0 S3.0
Without treatment (mV) -102.30 ± 2.14b -12.47 ± 0.75b -12.73 ± 0.25b
UP3 treatment (mV) -78.83 ± 1.76a -4.06 ± 0.41a -7.94 ± 0.17a
Lowercase letters show differences (p < 0.05) between the same formulation submitted or not to ultrasound treatment.
Tab. 3: Effect of process on the flow curves of biopolymers aqueous phase. Alginate 0.5% w/w (A), potato starch 3% w/w (S) and gelatin 1% w/w (G). Consistency index (k), behavior index (n) and ultrasonic probe 375 W/5 min (UP3). Formulations
k (mPa.sn)
n (-)
A0.5
207.7 ± 3.9b
0.6905 ± 0.0020a
A0.5 UP3
16.4 ± 2.4a
0.9377 ± 0.0223b
S3.0
3004.9 ± 95.6b
0.6017 ± 0.0049a
S3.0 UP3
2.7 ± 0.0a
1.0223 ± 0.0006b
G1.0
3.0 ± 0.9b
0.9673 ± 0.0372a
G1.0 UP3
0.6 ± 0.1a
1.1436 ± 0.0184b
Lowercase letters show differences (p < 0.05) between the same biopolymer solution submitted to different processes.
Tab. 4: Effect of gelatin concentration and process on the flow curves of biopolymer aqueous phase. Alginate 0.5 % (w/w), potato starch 3.0 % (w/w) and gelatin 0.1, 0.5, 1.0 and 1.5% (w/w). Consistency index (k), behavior index (n), rotor stator (RS), ultrasonic bath 150 W/5 min (UB), ultrasonic probe 150 W/5 min (UP1), ultrasonic probe 225 W/5 min (UP2), ultrasonic probe 375 W/5 min (UP3). Formulations SAG0.1 RS SAG0.1 UB SAG0.1 UP1 SAG0.1 UP2 SAG0.1 UP3 SAG0.5 RS SAG0.5 UB SAG0.5 UP1 SAG0.5 UP2 SAG0.5 UP3 SAG1.0 RS SAG1.0 UB SAG1.0 UP1 SAG1.0 UP2 SAG1.0 UP3 SAG1.5 RS SAG1.5 UB SAG1.5 UP1 SAG1.5 UP2 SAG1.5 UP3 SA RS SA UB SA UP1 SA UP2 SA UP3
k (Pa.sn) 3.92 ± 0.54 aB 3.18 ± 0.22 bA 0.14 ± 0.02 aA 0.05 ± 0.02 aA 0.02 ± 0.00 aA 6.12 ± 0.44 cB 4.00 ± 0.59 bB 0.17 ± 0.03 aA 0.04 ± 0.00 aA 0.02 ±0.00 aA 13.82 ± 0.78 cC 14.99 ± 0.55 dC 1.75 ± 0.81 bB 0.15 ± 0.03 aA 0.06 ± 0.01 aA 29.49 ± 0.57 cD 41.15 ± 0.57 dD 12.34 ± 0.53 bC 0.22 ± 0.06 aA 0.25 ±0.05 aA 3.61 ± 0.21 bA 3.99 ± 0.57 bB 0.17 ± 0.03 aA 0.04 ± 0.01 aA 0.02 ± 0.00 aA
n (-) 0.49 ± 0.01 aC 0.51 ± 0.01 aC 0.80 ± 0.01 bC 0.90 ± 0.04 cB 0.96 ± 0.01 dB 0.46 ± 0.01 aC 0.48 ± 0.02 aC 0.79 ± 0.02 bC 0.91 ± 0.01 cB 0.96 ± 0.02 dB 0.39 ± 0.00 aB 0.38 ± 0.00 aB 0.59 ± 0.07 bB 0.83 ± 0.02 cA 0.92 ± 0.01 dB 0.34 ± 0.00 aA 0.33 ± 0.00 aA 0.44 ± 0.00 bA 0.81 ± 0.02 cA 0.80 ± 0.02 cA 0.48 ± 0.01 aC 0.48 ± 0.02 aC 0.78 ± 0.02 bC 0.91 ± 0.02 cB 0.96 ± 0.01 dB
Lowercase letters show differences (p < 0.05) between the treatments for the same gelatin concentration. Uppercase letters shows differences (p < 0.05) between the different gelatin concentration, submitted to the same process.
Tab. 5: Polydispersity (SPAN), Sauter mean diameter (D3,2) and droplet size obtained by image analysis (ImageJ) at 0 and 7 days. Optical micrographs of the fresh emulsions (0 days) submitted to RS and UP3. Rotor stator (RS), ultrasonic bath at 150 W/5 min (UB), ultrasonic probe at 150 W/5 min (UP1), 225 W/5 min (UP2) and 375 W/5 min (UP3). Scale bar = 10 μm.
0 days 7 days 0 days RS 0 days UP3 SPAN D [3. 2] (µm) Size (µm) ImageJ SPAN D [3. 2] (µm) Size (µm) ImageJ G1.5 RS 1.52 ± 0.14 6.62 ± 0.32 dA 6.785 ± 1.235 d 1.86 ± 0.11 11.37 ± 0.60 dB 11.896 ± 1.689 d G1.5 UB 1.59 ± 0.01 6.03 ± 0.05 dA 5.589 ± 1.158 d 1.87 ± 0.11 8.56 ± 0.19 cB 8.123 ± 1.569 c cA c aB G1.5 UP1 2.58 ± 0.66 3.08 ± 0.06 3.125 ± 1.5981 2.17 ± 0.08 4.96 ± 0.40 4.874 ± 1.470 a bA b aB G1.5 UP2 1.29 ± 0.00 1.99 ± 0.38 2.026 ± 1.821 1.80 ± 0.01 4.26 ± 0.22 4.100 ± 1.339 a G1.5 UP3 1.45 ± 0.00 1.26 ± 0.01 aA 1.3690 ± 1.487 a 1.97 ± 0.31 6.56 ± 0.74 bB 6.649 ± 1.587 b aA a bB SAG0.1 RS 2.21 ± 0.07 4.38 ± 0.13 4.341 ± 1.609 1.90 ± 0.02 10.98 ± 0.29 4.262 ±1.483 a aA a bB SAG0.1 UB 2.24 ± 0.07 4.41 ± 0.13 4.539 ± 1.757 2.47 ± 2.22 9.45 ± 0.57 4.358 ± 2.099 a aA a aA SAG0.1 UP1 2.12 ± 0.11 4.26 ± 0.15 4.447 ± 1.489 2.21 ± 0.40 7.24 ± 0.37 3.934 ± 1.450 a SAG0.1 UP2 1.92 ± 0.01 4.70 ± 0.12 aA 4.566 ± 1.478 a 1.75 ± 0.09 9.42 ± 0.29 bB 5.536 ± 1.938 b bA b cB SAG0.1 UP3 1.66 ± 0.02 7.71 ± 0.25 5.382 ± 3.200 1.54 ± 0.11 13.91 ± 0.47 5.903 ± 2.454 b SAG0.5 RS 1.02 ± 1.05 2.72 ± 2.93 aA 3.645 ± 1.738 b 1.08 ± 1.02 5.21 ± 5.44 aA 4.085 ± 1.521 bc abA b aA SAG0.5 UB 1.62 ± 0.03 3.08 ± 0.13 3.694 ± 1.610 2.09 ± 0.11 4.69 ± 0.30 4.529 ± 1.464 c abA b aA SAG0.5 UP1 1.88 ± 0.06 3.72 ± 0.12 3.067 ± 1.812 2.86 ± 0.18 5.37 ± 1.44 3.947 ± 1.370 b abA a bB SAG0.5 UP2 1.77 ± 0.06 4.76 ± 0.26 1.926 ± 1.289 1.85 ± 0.24 8.67 ± 1.87 2.382 ± 1.022 ª SAG0.5 UP3 1.78 ± 0.07 4.86 ± 0.83 bA 1.522 ± 0.976 a 1.94 ± 0.10 7.59 ± 1.12 bA 2.556 ± 0.910 a abA d aA SAG1.0 RS 1.80 ± 0.02 3.01 ± 0.09 3.332 ± 1.384 2.09 ± 0.45 2.72 ± 0.28 4.289 ± 1.134 d aA cd bA SAG1.0 UB 1.64 ± 0.00 2.54 ± 0.00 3.041 ± 1.626 2.37 ± 030 4.55 ± 0.83 3.526 ± 1.365 c abA c bA SAG1.0 UP1 2.02 ± 0.04 3.44 ± 0.21 2.835 ± 1.334 2.55 ± 0.46 5.87 ± 1.37 3.368 ± 10127 c SAG1.0 UP2 1.67 ± 0.03 5.85 ± 1.02 cA 1.776 ± 0.911 b 1.82 ± 0.02 10.78 ± 1.52 cB 2.153 ± 1.089 b bcA cB SAG1.0 UP3 1.52 ± 0.06 4.64 ± 0.47 1.062 ± 0.62 ª 1.78 ± 0.04 9.35 ± 0.29 1.159 ± 0.259 ª SAG1.5 RS 1.89 ± 0.03 3.17 ± 0.04 aA 2.981 ± 1.100 b 1.62 ± 0.36 3.43 ± 0.68 aA 3.710 ± 1.304 b aA c aA SAG1.5 UB 1.68 ± 0.14 2.64 ± 0.14 3.508 ± 1.257 1.58 ± 0.27 3.11 ± 0.35 3.558 ± 0.909 b SAG1.5 UP1 2.45 ± 0.16 4.15 ± 0.75 abA 3.019 ± 1.107 b 3.96 ± 0.06 7.06 ± 3.49 bA 3.775 ± 1.097 b bcA cB SAG1.5 UP2 2.29 ± 0.27 5.54 ± 1.08 1.869 ± 1.146 ª 2.00 ± 0.07 9.89 ± 3.26 3.139 ± 1.106 b cA b bcA SAG1.5 UP3 1.70 ± 0.16 6.58 ± 0.62 2.370±1.476 1.89 ± 0.04 8.92 ± 3.42 2.210 ± 0.867ª aA aA SA RS 1.82 ± 0.18 10.72 ± 0.11 5.052±8.165ª 1.75 ± 0.30 14.68 ± 2.28 7.329 ± 13.631a aA abA SA UB 2.49 ± 0.81 12.85 ± 3.16 5.918±12.287ª 1.52 ± 0.10 18.01 ± 2.01 7.140 ± 13.713a aA abA SA UP1 1.57 ± 0.30 17.33 ± 0.39 5.407±4.306ª 1.44 ± 0.26 18.91 ± 3.80 7.569 ± 4.523a aA b bA SA UP2 1.18 ± 0.21 13.40 ± 0.79 21.649±7.544 1.35 ± 0.05 22.70 ± 1.25 20.375 ± 8.656b SA UP3 0.68 ± 0.01 33.38 ± 11.32 bA 33.995±11.095c 0.65 ± 0.02 58.83 ± 0.69 cB 37.733 ± 8.868b Lowercase letters show differences (p < 0.05) between the treatments for the same gelatin concentration for 0 and 7 days, uppercase letters show D3.2 differences (p < 0.05) between the 0 and 7 days for the same formulation. Formulations
S1350-4177(18)31159-3 https://doi.org/10.1016/j.ultsonch.2018.12.001 ULTSON 4398
To appear in:
Ultrasonics Sonochemistry
Received Date: Revised Date: Accepted Date:
1 August 2018 30 November 2018 1 December 2018
Please cite this article as: K.C.G. Silva, A.C.K. Sato, Sonication technique to produce emulsions: The impact of ultrasonic power and gelatin concentration, Ultrasonics Sonochemistry (2018), doi: https://doi.org/10.1016/ j.ultsonch.2018.12.001
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Sonication technique to produce emulsions: the impact of ultrasonic power and gelatin concentration Karen Cristina Guedes Silva a1, Ana Carla Kawazoe Satoa2. a
Department of Food Engineering, School of Food Engineering (FEA), University of
Campinas, CEP: 13083-862, Campinas, SP, Brazil. 1
[email protected], [email protected].
Corresponding authors: Department of Food Engineering, School of Food Engineering (FEA), University of Campinas. Rua Monteiro Lobato, 80, CEP: 13083-862, Campinas, SP, Brazil. Tel.: +55 19 35214088. E-mail addresses: [email protected] (K. C. G. Silva), [email protected] (A. C. K. Sato).
Highlights • Oil in water emulsions was prepared by rotor-stator and sonication processes. • Sonication process reduces the viscosity of the aqueous phase. • Higher ultrasonic power produced emulsions with higher stability. • Low protein concentration led to a coalescence during sonication. • Sonication was proposed as a suitable method for emulsion production.
ABSTRACT The production of food emulsions has increased the demand for processes, natural emulsifiers and stabilizers that provide reasonable stability. This study approaches the influence of parameters that affect the stability of emulsions produced by sonication, such as ultrasonic power (150, 225 and 375 W) and gelatin concentration, when producing alginate, potato starch and gelatin stabilized emulsions. The results showed that sonication reduced viscosity, surface charge and improved the interfacial properties of biopolymeric solutions. Emulsions presented visual kinetic stabilization after 7 days of storage. The increase of sonication power reduced particle size but increased flocculation. The use of ultrasonic power at 225 and 375W and gelatin in a concentration above 1.0 % resulted in stable emulsions with smaller particle size, which is desirable for its application in food systems.
Keywords: ultrasound, alginate, potato starch, emulsification, aqueous phase, stability.
1.
Introduction Emulsions are mixtures of immiscible liquids containing droplets of one liquid
(dispersed phase) dispersed in a continuous liquid phase. Emulsifying agents are the ones that provide stability to these systems. However, the emulsification process is entropically unfavorable, resulting in thermodynamically unstable systems [1]. Hence, it is important to develop highly stable food emulsions with no or minimal changes in its structure during its process and storage [2]. Different processes have been studied to achieve greater emulsion stability: high energy processes such as rotor-stator, ultrasound devices, high-pressure homogenizer, microfluidics, among others widely employed [3,4]. Previous studies have shown that, for high energy processes, the type of homogenization process, operating conditions (energy intensity, time and temperature), sample composition (oil and emulsifier concentrations), and physicochemical properties of the compounds (interfacial tension and viscosity), result in emulsions with different characteristics [5]. Commonly used to prepare pre-emulsions, rotor-stator process mixes liquids from separated phases, forcing them through a narrow gap between a rotating disc (rotor) and a static disk (stator). The rapid flow circulation and high shear within the narrow gap between the rotor and stator form droplets usually larger than those obtained
by other high energy processes [4,6]. It is possible to produce finer emulsions by sonication. In this process, the mechanism for droplet rupture is acoustic cavitation, which comprises the growth and collapse of micro gas bubbles dissolved in the medium, leading to droplets breakdown [7]. One can apply the ultrasonic devices to liquid media in the form of bath or probe. In ultrasonic baths, the transducers are evenly located at the bottom of the bath. However, interferences between waves can result in different wave intensity at different positions of the tank [8]. The probe system consists of a generator and a converter, which transforms energy into mechanical vibrations transmitted by a probe and directly applied to the sample afterwards [4]. In general, probe type sonicators have higher localized intensity and effect compared to those of bath type [9]. When compared to homogenizers and microfluidizers, sonication processes feature advantages such as being easier to clean and presenting a simpler system, not needing external pressure, using less energy and using less surfactant to obtain more homogenous emulsions [10,11]. With ultrasound application, benefits have been obtained in food processing and preservation, such as higher product yields, shorter processing times and lower costs, in addition to an environmentally friendly production [12]. Moreover, structural modifications from sonication of food structures enable the development of new products with unique properties and quality attributes [13]. In this sense, sonication can lead to functional modifications in polymer chains [14,15] and protein structures [16,17], by adding functional features, such as improvements in dissolution, hydration, hydrophobicity, emulsifying and rheological performances [16], besides improving antioxidant properties of certain food materials [18]. The ultrasound application has also shown to modify some physical properties of starch and biopolymers solutions, allowing its different applications [19–21]. Besides the process, biopolymers used to stabilize emulsions play a fundamental role in stabilization. Some papers in the literature have shown that the association of proteins and polysaccharides for emulsion production can increase stability during the process, compared to emulsions only stabilized by proteins [22]. This association contributes to the structure, texture and stability of food systems, due to the thickening or gelling behavior and surface properties of biopolymers [23,24]. Few studies provide a systematic approach to evaluate the effects of the ultrasound process on emulsion formation and stabilization, often associated with beneficial structural modifications of biopolymers. These modifications, allied with the
establishment of ideal concentrations of surfactant, structuring agents and energy employed in the process, may result in emulsions with improved characteristics and greater stability. Therefore, the objective of this study was to investigate the effects of the ultrasound process on the physical properties of natural biopolymers used to stabilize emulsions, such as protein and polysaccharides, and to understand how the ultrasound-promoted structural modifications of biopolymers can influence emulsion stability. Hence, it would be possible to apply these emulsions to food systems of varied characteristics as structuring agents, or even as encapsulation systems.
2. Materials and methods 2.1. Materials To prepare the emulsions, ingredients such as alginate FD 17X, donated by Danisco (France), A-type gelatin - bloom 280 (~5.5 isoelectric point and 87.92 g/100 ml protein content), supplied by Gelco (Brazil), potato starch from Zona Cerealista (Brazil) and sunflower oil, purchased from a local supermarket, were used. All other chemicals were of analytical grade or equivalent.
2.2. Methods 2.2.1. Aqueous phase preparation Solutions containing a mixture of 3% (w/w) potato starch, 0.5% (w/w) alginate and varying amounts of gelatin (0, 0.1, 0.5, 1.0 and 1.5 % (w/w)), named as SA, SAG0.1 ; SAG0.5 ; SAG1.0 and SAG1.5, respectively, were prepared. Solutions presented pH 6 ± 0.2. Starch and alginate were added into deionized water at room temperature, under constant magnetic stirring, until the alginate was dissolved and no lumps were observed. Subsequently, solutions were heated at 90 °C under constant magnetic stirring at 800 rpm, for 20 min. The solution was cooled to up to 45 °C before the addition of different concentrations of gelatin, maintaining constant magnetic stirring at 800 rpm for 20 min. Rheology of biopolymer solutions was evaluated. Pure starch (S), Alginate (A) and Gelatin (G) solutions were prepared at the same concentrations mentioned above, under the same conditions. These solutions were used as a control to evaluate the influence of ultrasound on the properties of each biopolymer separately.
2.2.2. Emulsion preparation
Oil-in-water emulsions were prepared by mixing 20% (w/w) sunflower oil and 80% (w/w) of aqueous phase solution, using a rotor-stator (RS) device (SilentCrusher M, Heidolph, Germany) at 20000 rpm for 4 min prior to bath or probe sonication. For the ultrasonic probe, an ultrasonic processor (QR 750 W, Ultronique, Brazil) with a frequency of 20 kHz, attached to a titanium probe (13 mm diameter) was used to sonicate the emulsions. Sonication time was fixed at 5 min, its power varying in 150 (UP1), 225 (UP2) and 375 W (UP3). A glass tube containing 30 mL of sample was introduced in jacketed beakers at 10 oC to dissipate the heat produced during sonication and keep the temperature below 45 oC. Ultrasonic bath (USC 1450, Unique, Brazil) was tested with a power of 150 W and 25 kHz frequency, and sonication time fixed at 5 min. Samples submitted to the ultrasonic bath treatment were named as UB. Emulsions were evaluated regarding their stability, microscopy and particle size. Sonication time was set at 5 min for bath and probe sonication equipment, once preliminary tests showed that increasing sonication time resulted in larger droplets (Table 1), indicating an over processing, possible due to a higher probability of droplet collision, resulting in coalescence [25].
2.3. Biopolymer solutions characterization 2.3.1. Differential Scanning Calorimetry Thermal properties of starch were analyzed using a scanning differential calorimeter (DSC 2920, TA Instruments, USA). The sample was heated at a 10 °C/min rate from 10 to 110 °C. The initial temperature (To), peak temperature (Tp), final temperature (Tf), and gelatinization enthalpy (ΔH) were obtained from sample thermograms analyzed using the Universal Analysis software (TA Instruments). 2.3.2. ζ - potential ζ-potential of biopolymers was determined using a Zetasizer Nano Series (Malvern Instruments, UK). Biopolymers solutions were measured with and without ultrasound application at the same concentration prepared for the aqueous phase.
2.3.3. Interfacial tension Interfacial tension between biopolymer solutions in water and sunflower oil was evaluated using a tensiometer (Tracker-S, Teclis, France) by the pendant (O/W) drop method. Interfacial properties of gelatin at different concentrations and the effects of
ultrasound on the interfacial properties of alginate, gelatin and starch was evaluated. The test was conducted for over 7200 s and the temperature was maintained at 25 °C throughout the test.
2.3.4. Rheological measurements Rheological properties of aqueous phase were characterized using a rheometer (AR1500ex, TA Instruments, USA) equipped with a cone/plate geometry (cone diameter: 40 mm; angle: 1,58° and gap size: 53 μm). Flow curves were obtained in the shear rate range of 0-1000 s-1 at 25 °C. Experimental data were described by the power law equation (1): ( 1) where and
is shear stress (Pa),
is consistency index (Pa. sn),
is shear rate (s-1)
is the flow behavior index (-).
2.4. Emulsions evaluation 2.4.1. Stability Immediately after preparation, 25 ml of each emulsion was poured into a cylindrical glass tube, sealed with a plastic cap and stored in BOD (MA 415 UR, Marconi, Brazil) at 25 °C for 7 days. The O/W emulsion stability was measured by the height of the bottom phase (Hb) and expressed according to creaming index (CI) (equation 2). ( (2) where H0 represents the initial height of emulsion. The analyses were carried out in duplicate.
2.4.2. Size distribution Measurement of droplet size distribution was performed in a Malvern Mastersizer S (Malvern Instruments, UK) equipped with accessory Hydro 2000S (Malvern Instruments, UK). Samples were dispersed into distilled water. Values of Sauter mean diameter, which is inversely proportional to the specific surface area of
droplets, were obtained (Equation 3). The polydispersity (Span) was also evaluated (Equation 4). n
D3, 2
n.d
3 i
n.d
2 i
i 1 n
i 1
( 3)
( 4) where ni is number of droplets with a diameter di, and
(µm) are
diameters at 10, 50 and 90% of cumulative volume, respectively.
2.4.3. Optical microscopy Emulsion microstructure was observed in an optical microscope (Axio Scope.A1, Carl Zeiss, Germany) with a 100× oil immersion objective lens. Images were captured with the software AxioVision Rel. 4.8 (Carl Zeiss, Germany). Optical microscopy was performed in freshly prepared emulsions and after 7 days of storage. The public domain software Image J v1.36b (http://rsb.info.nih.gov/ij/) was used to count droplets and measure their size.
2.6. Statistical analyses All trials were replicated three times and, unless otherwise stated, all analyses were carried out in two independent replicates. SISVAR software - Computer Statistical Analysis System [26] was used for variance analysis (ANOVA). Significant differences (p<0.05) between samples were determined by the Tukey procedure.
3. Results 3.1. Effect of gelatin concentration and process at the aqueous phase Aiming to evaluate the role of protein at different concentrations of emulsion stabilization, in addition to the effects of mechanical and sonication process in the structure of the isolated biopolymers, surface charge analysis, interfacial tension and rheology were performed for aqueous phase.
3.1.1. Differential Scanning Calorimetry
To determine the gelatinization temperature of the starch, we measured its thermal properties: T0 = 62.29 ± 0.11 °C, Tp = 65.64 ± 0.38 °C, Tf = 70.12 ± 0.11 °C and gelatinization enthalpy (∆H) (8.93 ± 1.35 J/g dry starch). The starch dilutions employed in this study were higher than 10 g water/g potato starch for all formulations, which ensures complete gelatinization, as the gelatinization temperature of the experiments was set at 90 °C/20 min. Evans & Hais [27] reported values for initial (~60 °C) and final (~70 °C) gelatinization temperature as a function of water content, for a ratio higher than 2 g water/g potato starch. 3.1.2. ζ - potential Zeta potential results of biopolymers showed that the sonication process (UP3) significantly reduced surface charge for all biopolymers (p<0.05) (Tab. 2).
3.1.3. Interfacial tension The evaluation of interfacial tension was carried out with the objective to characterize the interfacial properties of solutions with different gelatin concentrations and to investigate the influence of the sonication process on the emulsifying properties of biopolymers. Regarding interfacial properties of the protein, the increase of gelatin concentration exerts little influence on initial interfacial tension (7.2 – 7.8 mN/m). The difference became more pronounced at 7200 s, presenting results that varied between 3.1 to 4.3 mN/m for G1.5 to G0.1, respectively (Fig. 1A), which may be related to faster adsorption of gelatin at the droplet interface. Higher adsorption of the surfactant at the interface is associated with a reduction in interfacial properties, indicating that the surfactant and its amount have been effective, forming a protective barrier that prevents flocculation and retards destabilization [3]. However, it should be noted that even though higher concentration of surfactant reduces interfacial tension, the equilibrium in interfacial tension was not reached after 7200 s. Concerning the influence of the ultrasonic process (UP3), a decrease in the interfacial tension for gelatin and alginate solutions was observed compared to the same solutions without treatment. One can associate this result with the exposure of hydrophobic residues of the surfactant during sonication [28], which is also possible to observe from surface charge reduction after ultrasound application (Tab. 2). In contrast,
starch solution increased in interfacial tension when submitted to sonication process (UP3) (Fig. 1B).
3.1.4. Rheological measurements The ultrasound process employs a large amount of energy generating hydrodynamic shear forces, causing cavitation and collapse of gas bubbles. This phenomenon can change physical or physicochemical structures of the medium [16,29], which may result in the breaking of some chains, allowing greater fluidity of the continuous phase. Rheology results for isolated biopolymer solutions showed that sonication process (UP3) reduced the viscosity of alginate, gelatin and starch, which was of about two orders of magnitude for the latter (Tab. 3). Sonication can accelerate the depolymerization of starch gels, separating amylose and amylopectin units. In this situation, amylose units remain in the gel amorphous phase, which leads to the weakening of biopolymer network, thus reducing viscosity [19]. However, authors point out that despite sonication dramatically affecting solution viscosity, the chemical structure of polymer chains may not change [19]. The effect of process on the rheological properties of aqueous phase containing the associated biopolymers (Tab. 4) showed that rheological parameters of solutions submitted to probe sonication (UP) differed significantly (p<0.05) from the same formulation submitted to bath sonication (UB) or rotor stator (RS). The increase in sonication power reduced shear thinning of SAG solutions (n closer to 1) and reduced viscosity (associated to lower k) (Tab. 4). This effect may be due to possible breakdown of polymer chains caused by the sonication process [29,30]. Such breakdown reduces entanglements and consequently modifies intermolecular interactions between polymer molecules, reducing viscosity and leading to an approximation of Newtonian behavior [31,32]. Similar results regarding viscosity reduction with ultrasound application in polymers, such as glucomannan, pectin, proteins and starches from different sources have been reported in literature [19,32,33]. By comparing the same treatment through varying concentrations of gelatin, we observed that the increase in gelatin concentration increased viscosity for RS, UB and UP1. The effect of gelatin concentration is suppressed at higher powers (UP2 and UP3), with no significant differences (p>0.05) in viscosity (Tab. 4).
3.2. Effect of gelatin concentration and process on emulsions The power used during emulsification is a key point for the production of stable emulsions. At this stage of the work, we used different concentrations of gelatin when producing emulsions by stator rotor, ultrasonic bath and ultrasonic probe, at three different sonication powers. Stability, droplet size, and emulsion morphology were investigated.
3.2.1. Creaming index Emulsions were evaluated regarding stability by creaming index. Gelatin control emulsions submitted to RS, UB and UP1 process showed creaming index of 39.23 ± 3.39, 38.80 ± 2.06 and 24.71 % ± 6.60, respectively. For the same formulation, UP2 and UP3 process did not present phase separation at 7 days (Fig. 2A). The higher stability presented by emulsions solely containing gelatin was only observed for conditions of higher sonication power. This result supports the hypothesis that higher sonication power favors the exposure of surfactant hydrophobic residues, as fat-binding ability depends on the degree of exposure of gelatin hydrophobic residues [21] (Tab. 2 and Fig. 1B). For control emulsions without gelatin (SA), all formulations presented phase separation. The creaming index for RS, UB, UP1, UP2 and UP3 were: 60.92 ± 2.58, 61.82 ± 1.29, 66.73 ± 4.49, 43.97 ± 7.19 and 65.25 % ± 1.84, respectively (Fig. 2B). Despite sonication changing zeta potential, rheology and interfacial tension of alginate and starch solutions (Tab. 2 and 3 and Fig. 1B), neither of them act as emulsifiers, only contributing to system viscosity [23]. Thus, no stable emulsion was produced solely with alginate and starch. Protein association with potato starch and alginate (SAG0.1 to SAG1.5) provided visually stable emulsions under all tested processes and concentrations without creaming (data not shown), which emphasizes the importance of interfacial activity promoted by gelatin and of the hydrodynamic stabilization [3] promoted by starch and alginate.
3.2.2. Droplet size and morphology during storage Emulsions were evaluated in relation to their droplet size, polydispersity (SPAN), optical microscopy and image analysis at day 0 and after 7 days of storage (Tab. 5). Results showed that, for the same process, gelatin concentration (0.5 to 1.5%
(m/m)) did not show significant differences (p>0.05) (statistical data not shown) for emulsion droplet size. In other words, increase in sonication power was more effective in emulsion size reduction compared to gelatin concentration. Comparing the different processes at a fixed protein concentration, it was possible to verify the treatment influence on droplet size and, consequently, on stability. For the lowest protein concentration (SAG0.1) and for control emulsion (SA), we observed no significant differences (p>0.05) between RS, UB and UP1 treatments at day 0. However, the treatment with the highest sonication power (UP3) resulted in larger particle size immediately after the process (day 0). Higher sonication powers (UP2 and UP3) expose the droplets to large amounts of energy, not only led to droplet rupture but also collision and, therefore, coalescence and flocculation [34]. Possibly, only adding 0.1% gelatin was not sufficient to cover and stabilize the droplet interface [35]. In addition, viscosity reduction due to the process (Tab. 4) may have aggravated destabilization under these conditions [36]. During the 7 days of storage, destabilization became more pronounced, as can be observed in D[3,2] results (Tab. 5). Addition of 0.5 and 1.0% of protein to the emulsion resulted in increase of D[3,2] with increasing ultrasound power. However, size results obtained by image analysis indicated a reduction, as we can see in micrographs (Tab. 5). Such divergence between image analysis and laser diffraction (D[3,2] result), reflects flocculation caused by the process. The smallest droplet sizes of SAG0.5 and SAG1.0 came from the combination of gelatin concentration and ultrasound treatment. At these concentrations, it would be possible to coat the droplets formed during emulsification efficiently. Moreover, the most intense cavitation process (UP2 and UP3) reduced the viscosity of aqueous phase, facilitating surfactant mobility to the droplet interface, avoiding coalescence and reducing polydispersity [3]. Analyzing the results obtained for SAG1.5 emulsion, it was observed that droplet size also tends to reduce as ultrasonic power increases. However, SAG1.5 presented higher flocculation and apparent gelation at 7 days of storage (Tab. 5), despite being stored at room temperature (25 °C).
4. Discussion The use of higher sonication powers led to structural modifications in the aqueous phase of the isolated biopolymers, which reduced biopolymer viscosity (Table 3), interfacial tension (Figure 1), and surface charge intensity (Table 2). For gelatin,
these results are associated to the exposure of hydrophobic protein residues to sonication, reducing charge intensity [16]. We can associated such exposure with improvements in stability and dehydration of amphiphilic biopolymers, which decreases the energy required for protein adsorption at the oil-water interface [30]. The breakage of electrostatic and hydrophobic interactions between protein molecules with ultrasound has facilitated the hydrolysis process of gelatin [37] and increased solubility and thermal stability of treated whey protein[17]. Structural modifications provided by sonication has also been used to regulate viscosity [38] and modify foaming properties of products [13,39]. Structural modifications of several biopolymers have also been reported through sonication in the literature. Hosseini et al., [15] also reported the ζ-potential reduction for β-lactoglobulin and κ-carrageenan complexes with sonication. The authors attributed this phenomenon to the reduction of biopolymer reactivity after sonication, related to heterogeneous sonochemical interactions and structural changes. These changes can include a large number of sonochemical reactions in polysaccharides, including glycosylation, acetalization, oxidation and carbon bonds, capable of modifying interactions between polysaccharides and reducing reactive sites [12]. In this study, the zeta potential reduction observed for alginate and starch may stem from these sonochemical reactions. Viscosity reduction may depend on mechanical, structural properties of biopolymers, on linear or random coil configurations, as discussed by Iida et al., [19] for pectin and glucomannan. For individual biopolymers, we observed different intensity on viscosity reduction depending on their structure. Potato starch, the compound with the highest molecular weight [40], also showed the highest viscosity reduction compared to gelatin, a protein with random coil structure [41]. It is known that viscosity of the continuous phase directly affects the stability of dispersions [7]. By applying the highest powers (UP2 and UP3), we observed higher viscosity reduction at the aqueous phase. Such reduction provides greater surfactant mobility to interface droplets, which quickly reduces interfacial tension [42], as observed in this study. Behrend and Schubert [7] also reported that the higher viscosity of the continuous phase delays the mobility and stabilization of surfactant at the interface. Some studies have shown that the association of proteins and polysaccharides in emulsion production may increase the stability when compared to emulsions only stabilized by proteins [22]. The action of hydrocolloids as structuring, thickening and
gelling agents at the aqueous phase [43] provides a rheological control mechanism that immobilizes the oil droplets in an entangled biopolymer network [44]. In processes applying lower sonication powers (RS, UB and UP1) due to the higher viscosity, it is possible to say that part of the gelatin adhered to the surface of the droplets while part was dispersed in the medium with alginate and starch. This configuration characterizes depletion stabilization, a mechanism in which free (nonadsorbed) polymers inhibits aggregation [45], and in addition to the hydrodynamic stabilization from high viscosity. This behavior is often found in mixtures of food biopolymers [46]. Sonication at higher powers (UP2 and UP3) reduce droplet size and increased flocculation, which
is clear when observing the divergence between D[3,2] data,
obtained by laser diffraction, and size, obtained by optical microscopy images (Tab. 5). Under these conditions, it is possible to consider that the sonicated protein in solution has migrated to the oil interface, reducing the protein/polysaccharide ratio in the medium. It is worth noting that the energy employed in the ultrasound process is not sufficient to break the peptide bonds, but only non-covalent associative interactions, which does not cause changes in the primary structure of proteins [16]. Increases in free polysaccharide solution favor depletion flocculation [47, 48]. In this study, the presence of non-adsorbed alginate and starch in the continuous phase, which has no interfacial properties caused an osmotic effect, reducing the amount of solvent between the droplets and promoting attraction between them. This phenomenon occurs when protein-coated droplets are dispersed in an aqueous solution of other biopolymer not compatible with the protein [49]. In this case, the repulsion between gelatin located at the interface with other biopolymers occurs because all of them present a negative surface charge under these study conditions (Tab. 2). The increase in the size of bovine gelatin chains after 7 days of storage was also reported, indicating a reorganization of proteins in sub-aggregates due to non-covalent (electrostatic and hydrophobic) interactions [30]. This reorganization may have been responsible for the increased flocculation observed for the emulsions obtained with the highest intensity processes (UP2 and UP3) after 7 days (Tab. 5). The role of ultrasound in protein functionality modifications, such as thermal, structural and morphological attributes [17], and nanoemulsion production functionality has been studied [16]. However, few works in the literature have studied the effects of ultrasound process on structural modifications of the separate biopolymers and its
effects when biopolymers are associated, trying to understand how these modifications contribute to stabilizing emulsions. Besides, there has been increasing interest within the food industry to replace synthetic emulsifiers by natural ones, as an alternative to consumer-friendly products. Consequently, the trend of using biopolymers (proteins, starches) to produce and stabilize emulsions is growing [50] and there is still need to further understand the functionality of these biopolymers, when applied to different emulsification process. The results of this study indicated that the type of process and its intensity caused structural modifications in biopolymers. These modifications influenced the characteristics of the aqueous phase and consequently the formation of emulsions, resulting in systems with different characteristics.
4. Conclusions This study showed that the type of process provided more emulsion modifications, resulting in a wider variety of emulsions when compared to gelatin concentration. Higher sonication power changed the polymer network, with more influence on starch. Generally, sonication process decreased zeta potential, interfacial tension and viscosity of the biopolymers solutions, resulting in stable emulsions with smaller droplet size. Emulsions SAG0.5 and SAG1.0 sonicated at 225 and 375 W presented desirable characteristics for food application. Besides using only natural compounds, this study demonstrated that producing emulsions by sonication employing higher powers (UP2 and UP3) is a simple process that promotes desirable structural modifications in biopolymers.
Acknowledgments The authors thank the access to equipment and assistance provided by the laboratory of Process Engineering, Department of Food Engineering, School of Food Engineering from the State University of Campinas co-funded by Fapesp (2007/580175; 2009/54137-1; 2011/06083-0; 2015/11984-7). Karen Cristina Guedes Silva thanks CAPES for the assistantship. The authors thank Espaço da Escrita – Pró-Reitoria de Pesquisa - UNICAMP - for the language services provided.
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Fig. 1: (A) Interfacial tension of sunflower oil in gelatin solution at concentrations 0.1 % (w/w), 0.5 % (w/w), 1.0 % (w/w) and 1.5 % (w/w). (B) Interfacial tension of sunflower oil in gelatin 1.0 % (w/w), alginate 0.5 % (w/w) and starch 3.0 % (w/w) solutions. Solutions without treatment and submitted to ultrasonic power at 375 W/5 min (UP3) using the pendant drop method. Fig. 2: Stability of control emulsions at 7 days, submitted to rotor stator (RS), ultrasonic bath at 150 W/5 min (UB), ultrasonic probe at 150 W/5 min (UP1), 225 W/5 min (UP2) and 375 W/5 min (UP3). (A) Emulsion of sunflower oil in gelatin solution 1.5% (w/w). (B) Emulsion of sunflower oil in starch 3.0% (w/w) and alginate 0.5% (w/w) solutions.
A
B
G RS
G UB
G UP1
G UP2
G UP3
SA RS
SA UB
SA UP1
SA UP2
SA UP3
Tab. 1: Polydispersity (Span) and Sauter mean diameter (D3,2) of the SAG1.0 emulsion (0 days) submitted to ultrasonic bath (UB) and ultrasonic probe at 375 W (UP3) at 5, 10 and 15 min. Formulations SAG1.0 UB 5min SAG1.0 UB 10min SAG1.0 UB 15min SAG1.0 UP3 5min SAG1.0 UP3 10min SAG1.0 UP3 15min
Span 1.637 ± 0.002b 1.673 ± 0.012b 1.865 ± 0.008c 1.484 ± 0.010a 1.925 ± 0.005d 1.923 ± 0.012d
Lowercase letters show differences (p < 0.05) between the formulations
D [3, 2] (µm) 2.547 ± 0.002a 2.399 ± 0.028a 2.762 ± 0.077b 4.216 ± 0.140c 5.541 ± 0.082d 6.054 ± 0.040e
Tab. 2: Zeta potential of biopolymers alginate (A), gelatin (G) and potato starch (S) at pH 6 ± 0.2. Solutions without treatment and submitted to ultrasonic process at 375 W/5 min (UP3). Formulations A0.5 G1.0 S3.0
Without treatment (mV) -102.30 ± 2.14b -12.47 ± 0.75b -12.73 ± 0.25b
UP3 treatment (mV) -78.83 ± 1.76a -4.06 ± 0.41a -7.94 ± 0.17a
Lowercase letters show differences (p < 0.05) between the same formulation submitted or not to ultrasound treatment.
Tab. 3: Effect of process on the flow curves of biopolymers aqueous phase. Alginate 0.5% w/w (A), potato starch 3% w/w (S) and gelatin 1% w/w (G). Consistency index (k), behavior index (n) and ultrasonic probe 375 W/5 min (UP3). Formulations
k (mPa.sn)
n (-)
A0.5
207.7 ± 3.9b
0.6905 ± 0.0020a
A0.5 UP3
16.4 ± 2.4a
0.9377 ± 0.0223b
S3.0
3004.9 ± 95.6b
0.6017 ± 0.0049a
S3.0 UP3
2.7 ± 0.0a
1.0223 ± 0.0006b
G1.0
3.0 ± 0.9b
0.9673 ± 0.0372a
G1.0 UP3
0.6 ± 0.1a
1.1436 ± 0.0184b
Lowercase letters show differences (p < 0.05) between the same biopolymer solution submitted to different processes.
Tab. 4: Effect of gelatin concentration and process on the flow curves of biopolymer aqueous phase. Alginate 0.5 % (w/w), potato starch 3.0 % (w/w) and gelatin 0.1, 0.5, 1.0 and 1.5% (w/w). Consistency index (k), behavior index (n), rotor stator (RS), ultrasonic bath 150 W/5 min (UB), ultrasonic probe 150 W/5 min (UP1), ultrasonic probe 225 W/5 min (UP2), ultrasonic probe 375 W/5 min (UP3). Formulations SAG0.1 RS SAG0.1 UB SAG0.1 UP1 SAG0.1 UP2 SAG0.1 UP3 SAG0.5 RS SAG0.5 UB SAG0.5 UP1 SAG0.5 UP2 SAG0.5 UP3 SAG1.0 RS SAG1.0 UB SAG1.0 UP1 SAG1.0 UP2 SAG1.0 UP3 SAG1.5 RS SAG1.5 UB SAG1.5 UP1 SAG1.5 UP2 SAG1.5 UP3 SA RS SA UB SA UP1 SA UP2 SA UP3
k (Pa.sn) 3.92 ± 0.54 aB 3.18 ± 0.22 bA 0.14 ± 0.02 aA 0.05 ± 0.02 aA 0.02 ± 0.00 aA 6.12 ± 0.44 cB 4.00 ± 0.59 bB 0.17 ± 0.03 aA 0.04 ± 0.00 aA 0.02 ±0.00 aA 13.82 ± 0.78 cC 14.99 ± 0.55 dC 1.75 ± 0.81 bB 0.15 ± 0.03 aA 0.06 ± 0.01 aA 29.49 ± 0.57 cD 41.15 ± 0.57 dD 12.34 ± 0.53 bC 0.22 ± 0.06 aA 0.25 ±0.05 aA 3.61 ± 0.21 bA 3.99 ± 0.57 bB 0.17 ± 0.03 aA 0.04 ± 0.01 aA 0.02 ± 0.00 aA
n (-) 0.49 ± 0.01 aC 0.51 ± 0.01 aC 0.80 ± 0.01 bC 0.90 ± 0.04 cB 0.96 ± 0.01 dB 0.46 ± 0.01 aC 0.48 ± 0.02 aC 0.79 ± 0.02 bC 0.91 ± 0.01 cB 0.96 ± 0.02 dB 0.39 ± 0.00 aB 0.38 ± 0.00 aB 0.59 ± 0.07 bB 0.83 ± 0.02 cA 0.92 ± 0.01 dB 0.34 ± 0.00 aA 0.33 ± 0.00 aA 0.44 ± 0.00 bA 0.81 ± 0.02 cA 0.80 ± 0.02 cA 0.48 ± 0.01 aC 0.48 ± 0.02 aC 0.78 ± 0.02 bC 0.91 ± 0.02 cB 0.96 ± 0.01 dB
Lowercase letters show differences (p < 0.05) between the treatments for the same gelatin concentration. Uppercase letters shows differences (p < 0.05) between the different gelatin concentration, submitted to the same process.
Tab. 5: Polydispersity (SPAN), Sauter mean diameter (D3,2) and droplet size obtained by image analysis (ImageJ) at 0 and 7 days. Optical micrographs of the fresh emulsions (0 days) submitted to RS and UP3. Rotor stator (RS), ultrasonic bath at 150 W/5 min (UB), ultrasonic probe at 150 W/5 min (UP1), 225 W/5 min (UP2) and 375 W/5 min (UP3). Scale bar = 10 μm.
0 days 7 days 0 days RS 0 days UP3 SPAN D [3. 2] (µm) Size (µm) ImageJ SPAN D [3. 2] (µm) Size (µm) ImageJ G1.5 RS 1.52 ± 0.14 6.62 ± 0.32 dA 6.785 ± 1.235 d 1.86 ± 0.11 11.37 ± 0.60 dB 11.896 ± 1.689 d G1.5 UB 1.59 ± 0.01 6.03 ± 0.05 dA 5.589 ± 1.158 d 1.87 ± 0.11 8.56 ± 0.19 cB 8.123 ± 1.569 c cA c aB G1.5 UP1 2.58 ± 0.66 3.08 ± 0.06 3.125 ± 1.5981 2.17 ± 0.08 4.96 ± 0.40 4.874 ± 1.470 a bA b aB G1.5 UP2 1.29 ± 0.00 1.99 ± 0.38 2.026 ± 1.821 1.80 ± 0.01 4.26 ± 0.22 4.100 ± 1.339 a G1.5 UP3 1.45 ± 0.00 1.26 ± 0.01 aA 1.3690 ± 1.487 a 1.97 ± 0.31 6.56 ± 0.74 bB 6.649 ± 1.587 b aA a bB SAG0.1 RS 2.21 ± 0.07 4.38 ± 0.13 4.341 ± 1.609 1.90 ± 0.02 10.98 ± 0.29 4.262 ±1.483 a aA a bB SAG0.1 UB 2.24 ± 0.07 4.41 ± 0.13 4.539 ± 1.757 2.47 ± 2.22 9.45 ± 0.57 4.358 ± 2.099 a aA a aA SAG0.1 UP1 2.12 ± 0.11 4.26 ± 0.15 4.447 ± 1.489 2.21 ± 0.40 7.24 ± 0.37 3.934 ± 1.450 a SAG0.1 UP2 1.92 ± 0.01 4.70 ± 0.12 aA 4.566 ± 1.478 a 1.75 ± 0.09 9.42 ± 0.29 bB 5.536 ± 1.938 b bA b cB SAG0.1 UP3 1.66 ± 0.02 7.71 ± 0.25 5.382 ± 3.200 1.54 ± 0.11 13.91 ± 0.47 5.903 ± 2.454 b SAG0.5 RS 1.02 ± 1.05 2.72 ± 2.93 aA 3.645 ± 1.738 b 1.08 ± 1.02 5.21 ± 5.44 aA 4.085 ± 1.521 bc abA b aA SAG0.5 UB 1.62 ± 0.03 3.08 ± 0.13 3.694 ± 1.610 2.09 ± 0.11 4.69 ± 0.30 4.529 ± 1.464 c abA b aA SAG0.5 UP1 1.88 ± 0.06 3.72 ± 0.12 3.067 ± 1.812 2.86 ± 0.18 5.37 ± 1.44 3.947 ± 1.370 b abA a bB SAG0.5 UP2 1.77 ± 0.06 4.76 ± 0.26 1.926 ± 1.289 1.85 ± 0.24 8.67 ± 1.87 2.382 ± 1.022 ª SAG0.5 UP3 1.78 ± 0.07 4.86 ± 0.83 bA 1.522 ± 0.976 a 1.94 ± 0.10 7.59 ± 1.12 bA 2.556 ± 0.910 a abA d aA SAG1.0 RS 1.80 ± 0.02 3.01 ± 0.09 3.332 ± 1.384 2.09 ± 0.45 2.72 ± 0.28 4.289 ± 1.134 d aA cd bA SAG1.0 UB 1.64 ± 0.00 2.54 ± 0.00 3.041 ± 1.626 2.37 ± 030 4.55 ± 0.83 3.526 ± 1.365 c abA c bA SAG1.0 UP1 2.02 ± 0.04 3.44 ± 0.21 2.835 ± 1.334 2.55 ± 0.46 5.87 ± 1.37 3.368 ± 10127 c SAG1.0 UP2 1.67 ± 0.03 5.85 ± 1.02 cA 1.776 ± 0.911 b 1.82 ± 0.02 10.78 ± 1.52 cB 2.153 ± 1.089 b bcA cB SAG1.0 UP3 1.52 ± 0.06 4.64 ± 0.47 1.062 ± 0.62 ª 1.78 ± 0.04 9.35 ± 0.29 1.159 ± 0.259 ª SAG1.5 RS 1.89 ± 0.03 3.17 ± 0.04 aA 2.981 ± 1.100 b 1.62 ± 0.36 3.43 ± 0.68 aA 3.710 ± 1.304 b aA c aA SAG1.5 UB 1.68 ± 0.14 2.64 ± 0.14 3.508 ± 1.257 1.58 ± 0.27 3.11 ± 0.35 3.558 ± 0.909 b SAG1.5 UP1 2.45 ± 0.16 4.15 ± 0.75 abA 3.019 ± 1.107 b 3.96 ± 0.06 7.06 ± 3.49 bA 3.775 ± 1.097 b bcA cB SAG1.5 UP2 2.29 ± 0.27 5.54 ± 1.08 1.869 ± 1.146 ª 2.00 ± 0.07 9.89 ± 3.26 3.139 ± 1.106 b cA b bcA SAG1.5 UP3 1.70 ± 0.16 6.58 ± 0.62 2.370±1.476 1.89 ± 0.04 8.92 ± 3.42 2.210 ± 0.867ª aA aA SA RS 1.82 ± 0.18 10.72 ± 0.11 5.052±8.165ª 1.75 ± 0.30 14.68 ± 2.28 7.329 ± 13.631a aA abA SA UB 2.49 ± 0.81 12.85 ± 3.16 5.918±12.287ª 1.52 ± 0.10 18.01 ± 2.01 7.140 ± 13.713a aA abA SA UP1 1.57 ± 0.30 17.33 ± 0.39 5.407±4.306ª 1.44 ± 0.26 18.91 ± 3.80 7.569 ± 4.523a aA b bA SA UP2 1.18 ± 0.21 13.40 ± 0.79 21.649±7.544 1.35 ± 0.05 22.70 ± 1.25 20.375 ± 8.656b SA UP3 0.68 ± 0.01 33.38 ± 11.32 bA 33.995±11.095c 0.65 ± 0.02 58.83 ± 0.69 cB 37.733 ± 8.868b Lowercase letters show differences (p < 0.05) between the treatments for the same gelatin concentration for 0 and 7 days, uppercase letters show D3.2 differences (p < 0.05) between the 0 and 7 days for the same formulation. Formulations